Microscopy ListServer Archive Output

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Hi Isha,

here is the permalink to MIAWiki page on the topic:
http://confocal-manawatu.pbworks.com/w/page/44308564/Color%20SEM%20Images

Hope that will be helpful.

Cheers,
Dmitry
MIAWiki for Mass Collaboration

On Mon, January 2, 2012 5:57 am,
microscopylistserver-noreply-at-microscopy.com wrote:
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} Email: isha.mutreja-at-gmail.com
} Name: isha mutreja
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} Organization: Ulster University
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} Title-Subject: [Filtered] Biomedical Sciences
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} Message: How to process SEM images by adobe photoshop to add coloured
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} 8, 25 -- From microscopylistserver-noreply-at-microscopy.com Sun Jan 1
} 10:51:40 2012
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--
Dr. Dmitry Sokolov
Institute of Fundamental Sciences
Massey University, Palmerston North
New Zealand

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Contents Retrieved from Microscopy Listserver Archives
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Dear Nestor,

A very Happy New Year to you, too! I would like to thank you for the
Fantastic job you do with the listserver. It has been a very positive
influence in many lives, and careers.

Best Regards,
Darrell Miles

microscopylistserver-noreply-at-microscopy.com wrote on 01/01/2012 09:03:45
AM:

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|--br--||--font size=2 face="sans-serif"--|Dear Nestor,|--/font--|
|--br--|
|--br--||--font size=2 face="sans-serif"--|A very Happy New Year to you,
too! I
would like to thank you for the Fantastic job you do with the listserver.
It has been a very positive influence in many lives, and
careers.|--/font--|
|--br--|
|--br--||--font size=2 face="sans-serif"--|Best Regards,|--/font--|
|--br--||--font size=2 face="sans-serif"--|Darrell Miles|--/font--|
|--br--|
|--br--|
|--br--|
|--br--||--tt--||--font
size=2--|microscopylistserver-noreply-at-microscopy.com wrote
on 01/01/2012 09:03:45 AM:|--br--|
|--br--|
> [image removed] |--/font--||--/tt--|
|--br--||--tt--||--font size=2--|> |--br--|
> [Microscopy] Administrivia: Happy New Year - 2012|--/font--||--/tt--|
|--br--||--tt--||--font size=2--|> |--br--|
> microscopylistserver-noreply |--/font--||--/tt--|
|--br--||--tt--||--font size=2--|> |--br--|
> to:|--/font--||--/tt--|
|--br--||--tt--||--font size=2--|> |--br--|
> Darrell Miles|--/font--||--/tt--|
|--br--||--tt--||--font size=2--|> |--br--|
> 01/01/2012 09:04 AM|--/font--||--/tt--|
|--br--||--tt--||--font size=2--|> |--br--|
> Please respond to microscopylistserver-noreply|--/font--||--/tt--|
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> |--br--|
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> Happy New Year Colleagues;|--br--|
> |--br--|
> Welcome to the 20th year of operation of the Microscopy
ListServer|--br--|
> a free service to the world wide microscopy community,
sponsored|--br--|
> jointly by your Friendly Neighborhood SysOp and the Microscopy
Society|--br--|
> of America.|--br--|
> |--br--|
> During 2011, the ListServer delivered 2471 messages
to
over 3300|--br--|
> subscribers|--br--|
> around the world, with minimal hassels (that I know
about).
For |--br--|
> those of you that are|--br--|
> statistics junkies this year you generated 275+ Gb of Email
traffic and over|--br--|
> 8.1 Million Email messages were sent out this year by my tired little
server.|--br--|
> As usual you don't want to know how much Junk Mail and spam has
been|--br--|
> filtered out.|--br--|
> |--br--|
> |--br--|
> The complete Microscopy ListServer Archives for 2011-1993 (~ are
on-line
at|--br--|
> |--br--|
> |--/font--||--/tt--||--a
href=http://www.microscopy.com/--||--tt--||--font
size=2--|http://www.microscopy.com|--/font--||--/tt--||--/a--||--tt--||--font
size=2--|.|--br--|
> |--br--|
> A couple of IMPORTANT reminders:|--br--|
> |--br--|
> If you leave on vacation/holiday use the on-line form to
UNSUBSCRIBE|--br--|
> your Email address from the listserver. The out-of-office /
on-vacation|--br--|
> autoreply messages are a real nuisance to posters.|--br--|
> |--br--|
> Do not reply to message with the return address of:|--br--|
> |--br--|
> MicroscopyListserver-noreply-at-microscopy.com|--br--|
> |--br--|
> these are messages forwarded usually from the WWW posting form. They
do not|--br--|
> go back to the poster but rather into a black hole, which I rarely
check.|--br--|
> If you see a message that has this "No-Reply" return
address
please post your|--br--|
> reply/comment/answer to:|--br--|
> |--br--|
> Microscopy-at-microscopy.com|--br--|
> |--br--|
> or if you wish to reply privately, look at the username in the body
of|--br--|
> the message the originators Email address is usually listed
therein.|--br--|
> |--br--|
> As always if you have questions about suitability of postings
or|--br--|
> are having problems, feel free to contact me at
(zaluzec-at-microscopy.com)|--br--|
> |--br--|
> Cheers,|--br--|
> |--br--|
> Nestor|--br--|
> Your Friendly Neighborhood SysOp|--br--|
> |--br--|
> ==============================Original
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Colleagues

I have approximatley 200 ml of 80 mM Ammonium Molybdate
solution left by a user in our Lab. The solution was used
as a stain for biological specimens. Can anyone familiar
with this stain point me to information concerning proper
method of disposal.

The MSDS sheet on Ammonium Molybdate indicates care should
be taken.

Thanks...

Nestor
Your Friendly Neighborhood SysOp.

--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory
Fellow of the Microscopy Society of America
E.P. Wigner Fellow - Oak Ridge National Laboratory
Senior Fellow of the Computational Institute - University of Chicago
Past President Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a Mac !

===========================================

==============================Original Headers==============================
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A postdoctoral position is available at the University of Wisconsin-Madison to work on electron tomography and cryo-electron microscopy of membrane deformation mechanisms mediated by ESCRT proteins. The postdoctoral researcher will work as part of a multidisciplinary team integrated by cell biologists, biophysicists, and virologists to understand structural aspects of ESCRT-mediated membrane budding in multiple organisms/systems, including plants, worms, and mammalian cultured cells.
�
The project entails imaging of plastic and vitreous sections of plant and animal tissues by electron tomography.

Applicants must have a Ph.D. and proven experience in the field of electron microscopy, electron tomography, and cryo-electron microscopy. Experience with vitreous sectioning and single particle analysis is desirable. Excellent written and oral skills and the ability to work collaboratively with others are required.

For more information of ongoing research in the labs involved in this project, please visit the following websites:
http://www.botany.wisc.edu/otegui/welcome.html
http://www.bmolchem.wisc.edu/faculty/audhya.html
http://www.mcardle.wisc.edu/faculty/bio/ahlquist_p.html

Interested candidates should submit a letter stating experience, and also include a CV and the names and contact information (phone and e-mail) of individuals who can provide a professional letter of references in a single pdf file to Dr Marisa Otegui (otegui-at-wisc.edu).

Marisa Otegui
Associate Professor
B119 Birge Hall
430 Lincoln Drive
Department of Botany
Madison, WI 53706
Lab webpage http://www.botany.wisc.edu/otegui/welcome.html

Phone: (608)265-5703
Fax: (608)262-7509

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Hi Isha,

As well as Dmitry's www5.pbrc.hawaii.edu/microangela etc links, below is
some info copy and pasted from a very similar discussion on colouring B&W
TEM images back in January 2010 on the microscopy list-server [to find all
these posts, the threads were under the title: 'Image Colorizing']:

I tried out colorizing an SEM image with Photoshop CS4 and the results
seemed fine:

Original SEM image of pollen
http://en.wikipedia.org/wiki/File:Misc_pollen.jpg

I Colorized the image using the method in my last post, see below [you could
use Photoshop CS4 or Elements 8]
http://www.well.ox.ac.uk/cytogenetics/pollen/pollen_colorized.jpg
It took well under an hour to select and colorize the whole image [and I
found it quite therapeutic].

I also applied a Pseudocolor look-up table [LUT] applied using Photoshop CS4
[and Elements 8 has similar tools]
http://www.well.ox.ac.uk/cytogenetics/pollen/pollen_pseudo.jpg
For details of creating pseudocolor [false colour] with LUTs [e.g. Image,
Mode, Color table] with Photoshop CS4 see the likes of
http://www.imagingandanalysis.com/pseudo.pdf
Adobe's help, and search the internet.

----------------------------------------------------------------------------
----------------------------------------------------------------------------
------------------
Details of the colouring technique in Photoshop CS4:

Hi Isha,

It is quite easy, if time consuming, to do this Photoshop CS4 or CS5 [CS5
Extended's only �130 with Educational discounts - on a really tight budget
the likes Serif PhotoPlus X4 or Adobe Elements 10 will suffice].

There's always different ways to approach something like this in Photoshop
CS5 [or earlier versions], but I'd try:

Load the SEM Photo and manually select the region you wish to colourize,
using say the 'quick selection tool' - and you can add/reject [shift/alt]
bits of selected regions with a lasso tool. Then ensure the image is
converted to RGB colour [image, mode]. I'd then go to 'image, adjustments,
colour balance' and lightly adjust the colour balance of the selected region
and then "you can be brown, you can be blue, you can be violet sky", e.g.
move the colour balance slider from cyan to red and the selected object with
become progressively more red, with the underlying structural details
intact.
Then work through the entire image. Select all similar objects [to be the
same colour] within the image as one multiple region for colourizing [hold
shift as you select]. Make a mistake and use Edit, Undo. You can say
slightly adjust contrast and brightness, or curves, or shadow/highlights
within those selected regions as well.

You don't really need layers, you could work only on the main image
[background] bit by bit - save regularly under new file names as Photoshops
Undo [step backwards] is limited if, after a lot of work, you don't like the
way the image is turning out.

You could do all this with Photoshop Elements 8 as well [�37 with
discounts]. Do all the above Photoshop stuff using the similar Elements
selection tools, then [instead of 'colour balance'] go to: Enhance, Adjust
Colour, Adjust Hue/Saturation [and make sure the 'Colorize' box is ticked].

It will take a while to manually edit the entire image ['View, Zoom' to aid
tracing], but I doubt any image processing software could fully
'automatically' select the regions you want to colorize, particularly with a
greyscale SEM/TEM image. Photoshop's 'Quick selection tool' will have a go
though [the tools selection effect is adjustable in the upper menu bar].

I doubt the likes of alternative 'Pseudocolour' LUT effects will provide the
subtle colouring you require, although they may work adequately on TEM
images of sections.

Hope this helps,

Regards

Keith

---------------------------------------------------------------------------
Dr Keith J. Morris,
Molecular Cytogenetics and Microscopy Core,
Laboratory 00/069 and 00/070,
The Wellcome Trust Centre for Human Genetics,
Roosevelt Drive,
Oxford� OX3 7BN,
United Kingdom.

Telephone:� +44 (0)1865 287568
Email:� kjmorris-at-well.ox.ac.uk
Web-pages: http://www.well.ox.ac.uk/molecular-cytogenetics-and-microscopy

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Email: isha.mutreja-at-gmail.com
Name: isha mutreja

Organization: Ulster University

Title-Subject: [Filtered] Biomedical Sciences

Message: How to process SEM images by adobe photoshop to add coloured
effects to those images. Thank
you in anticipation. Looking forward for some help and guidance.

Login Host: 2.220.237.78
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Hi, Isha-

I tried replying in the thread, but the spam filter thought I had an
attachment and refused it, so Keith's instructions are not "below".

*************************************************************************
Yes, follow Keith's instructions below. Learn to use your selection tools
well, deciding, for example, if you want to feather the selection so it
looks more natural. I do like to use Selection - Save Selection so I can
save each selection as I go (and not lose all that hard work) and go back
and play with them later. Selectin the background first is often easiest,
and then you can select the inverse. You can use the Brush tool, the paint
bucket, or Edit - Fill, but in each case remember to select Color mode
instead of Normal (up on the menu bar). It is not automatic; it is time
consuming, but it can also be very therapeutic! These days, instead of
maintaining my MicroAngela website I make enamel jewelry under a
microscope... Same idea; surround an area with wire and then fill in with
colors. Yes, I did like coloring books when I was a child!

Aloha,
Tina
http://www5.pbrc.hawaii.edu/microangela/

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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Dear Listers
Thanks in advance for your time. A researcher here (see below) would like to quench her GFP signal. Any thoughts on how best to quench GFP with out disrupting the other antigens?

"Another question I have is if you known how to quench a GFP signal? I've been researching online and asking people and it seems that there isn't a good consensus on how to do it. I've heard that acetone fixation will work but I'm afraid that it might mess up my other antigens. I can't re-clone my virus, so I'm kind of stuck with the GFP...."

Happy New Year
Mary

Microscopy Facility Director
Neuroscience Research Institute&
Molecular, Cellular& Developmental Biology
Biology 2 Building, room 5173
The University of California
Santa Barbara, CA 93106-5060

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Recently I've been asked a question about upper magnification range with the SEM. I know upper magnification is going to depend on each SEM, type of filament, skill of the operator, sample type, definitions of what's useful and other factors I haven't even thought of. But, as a general rule of thumb, what's the normal upper magnification with meaningful resolution (Definition: I can clearly see the features I want and would publish the image) for a ordinary hair pin filament SEM?

any thoughts?

Thanks!!!!

Frank Karl
Microscopist
ARDL
2887 Gilchrist Road
Akron, Ohio 44305
330-794-6600

________________________________
This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Sales Representative - Nanotechnology

The Company:

Our client is a leading developer and producer of innovative high-tech
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the market leaders in each of the fields of Microscopy, Confocal Laser
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The Opportunity:

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Other: Company Car - Pension Plan - Home Office - Laptop - Mobile Phone -
Internet paid by company

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Personify
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Hi, Frank,

Having taken about 20,000 biological SEM photos with a tungsten filament SEM, the useful top magnification for images of the surface of cells runs about 20,000x. You can push this to as much as 100,000x for organelles of ideally prepared samples if your instrument can go to 35-40kV and you are working with gold-palladium coated samples. I've imaged viruses budding from the surface of cells at 20-40kX before by SEM and had good photos. With material samples such as metals, where you have less beam penetration and more signal, you will get better signal to noise ratio and better high-end magnification photos, so 100,000x photos are not much of a problem. My microscope maxed out at 180,000x, and I could get grainy photos of my resolution sample at that magnification, but could still resolve 5nm on a tin ball on carbon sample.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
________________________________________
X-from: frank_karl-at-ardl.com [frank_karl-at-ardl.com]
Sent: Friday, January 06, 2012 9:08 AM
To: Haller, Edward

Recently I've been asked a question about upper magnification range with the SEM. I know upper magnification is going to depend on each SEM, type of filament, skill of the operator, sample type, definitions of what's useful and other factors I haven't even thought of. But, as a general rule of thumb, what's the normal upper magnification with meaningful resolution (Definition: I can clearly see the features I want and would publish the image) for a ordinary hair pin filament SEM?

any thoughts?

Thanks!!!!

Frank Karl
Microscopist
ARDL
2887 Gilchrist Road
Akron, Ohio 44305
330-794-6600

________________________________
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As with the SEM of Ed Haller, our 28 y/o Hitachi S570 gave reasonably
good images up to 100KX. I believe this is probably a good expectation
for most tungsten-filament based SEMs. Some might give even better,
while others (like entry level or variable pressure instruments) may
not perform as well.

Of course, trimming the SEM (alignment, stigmation, good vacuum,
stable and conductive specimen) will have a tremendous effect on the
final image.

Cheers,
--
John J. Bozzola, Ph.D., Professor & Director of IMAGE (Retired : -)
Integrated Microscopy & Graphics Expertise
Southern Illinois University
750 Communications Drive
Carbondale, IL 62901
Phone: 618-453-3730

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Venu,

Look up this paper:

Rodighiero, S., et al., Fixation, mounting and sealing with nail polish of cell specimens lead to incorrect FRET measurements using acceptor photobleaching. Cellular Physiology and Biochemistry, 2008. 21(5-6): p. 489-498.

John Oreopoulos
Research Assistant
Spectral Applied Research
Richmond Hill, Ontario
Canada
www.spectral.ca

On 2012-01-09, at 3:15 PM, oshel1pe-at-cmich.edu wrote:

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} } Date: Mon, 9 Jan 2012 12:02:29 -0800
} } Reply-To: Venu Polineni {vens06-at-gmail.com}
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} } realname - Venu Polineni
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} } EDUCATION - Graduate College
} } SUBJECT_OF_QUESTION - FRET
} } QUESTION - Is there a specific chemical like saponin to intensify a
} } FRET signal? Or are there specific agents/detergents/fixatives that
} } either intensify or bleach a FRET signal.
} }
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I have a colleague at Creighton University who is trying to repair a
LKB-Pyramitome model 11800 and needs a "toothed" belt. Can anyone help?
She doesn't subscribe to this listserver so I'll pass along any
information to her. Thanks.

Tom Bargar
University of Nebraska Medical Center
Core Electron Microscopy Research Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu

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2, 23 -- Subject: spare parts for LKB-Pyramitome model 11800
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Hi Tom,

I am not familiar with the particular model, but if dimensions of the
belt can be measured or deducted from the design of system then good
places to start looking for it may be:

http://www.smallparts.com/b/16411421/ref=sp_iss_16411421

http://www.mcmaster.com/#timing-belts/=fre4ok

In unlikely case that McMaster and SmallParts would not have what you
need - try to Google for "timing belt", I am sure that there are plenty
of other places to get it.

Cheers :)
Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 1/11/2012 10:16 AM, tbargar-at-unmc.edu wrote:
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} I have a colleague at Creighton University who is trying to repair a
} LKB-Pyramitome model 11800 and needs a "toothed" belt. Can anyone help?
} She doesn't subscribe to this listserver so I'll pass along any
} information to her. Thanks.
}
} Tom Bargar
} University of Nebraska Medical Center
} Core Electron Microscopy Research Facility
} 986395 Nebraska Medical Center
} Omaha, NE 68198-6395
} 402-559-7347
} tbargar-at-unmc.edu
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--
I know that most EM labs have moved away from EM film although quite
a few labs in my field still use it. I just got an email from my
vendor who gave me an update on Kodak film. I am posting that
message in its entirety. I thought that a number of other users on
the list might like this information so I hope I am not crossing the
line into advertising.

Norm Olson
University of California San Diego
***********************
Norm-

Good Afternoon
Below is an email from my Kodak Product Manager about the Kodak
SO-163 EM film you purchase from us. I have had a number of calls
pertaining to the film in the last few days since Kodak announced
their probability of filing bankruptcy.
Please don't hesitate to call or email me if there are any additional
questions you may have. I have stock available if you need to place
an order.
Best Regards,
Sheila

Sheila Danahy
Sales Representative
Aremac Holdings
Tritech Forensics
Phone #866-972-6464 X 3128
Fax #866-682-0940
www.tritechforensics.com

KODAK Electron Microscope 4489 and KODAK Electron Image Film SO-163
are manufactured and sold by Carestream Molecular Imaging. You can
assure your customers that these products will be readily available
in the future as far as we can see.

Some background information. When Eastman Kodak Company spun off the
health sciences part of their business which became Carestream Health
in 2007, Carestream Health purchased all the film factories and
manufacturing equipment as well as retaining all the skilled staff to
continue producing the film products. Based on the value of the
Kodak name, Carestream has an agreement with Eastman Kodak Company to
sell the product under the Kodak name.

The film is made on the same machines by the same talented
manufacturing teams by Carestream Health. You can inform customers
you have spoken directly with the film source company and the EM film
is and will be available for them whenever they need it.

Please do not hesitate to contact me if I can be of assistance on any matter.

Best regards,
Eric

Eric Ambrose
Product Manager - Media Products
Carestream Molecular Imaging

eric.ambrose-at-carestream.com
P 585-627-8762
M 585-330-1646

Carestream Health, Inc.
150 Verona Street
Rochester, NY 14608

==============================Original Headers==============================
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If the belt can be measured, here are two sources to check:
http://www.smallparts.com
or
http://www.pic-design.com

Woody
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} I have a colleague at Creighton University who is trying to repair a
} LKB-Pyramitome model 11800 and needs a "toothed" belt. Can anyone help?
} She doesn't subscribe to this listserver so I'll pass along any
} information to her. Thanks.
}
}

--

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using the WWW based Form at http://www.microscopy.com/MLFormMail.html
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Email: mmh-at-umn.edu Name: Marc Hirschmann

Organization: University of Minnesota

Title-Subject: [Filtered] Job Opportunity: Lab Mgr. for electron microprobe lab

Message: Electron Microprobe Research Scientist, University of Minnesota, Minneapolis
The Department of Earth Sciences is seeking applications for an Electron Microprobe Research
Scientist. Responsibilities will include operation, supervision and routine maintenance of a
5-spectrometer JEOL 8900 electron microprobe. Experience analyzing geological samples by EMPA is
essential. Some experience maintaining a microprobe is desirable, but some amount of on-the-job
training is expected and the instrument continues to be on a JEOL service contract. The successful
applicant will also be expected to provide user training, including teaching a course in microprobe
theory and applications, and to consult/collaborate with University researchers and students on
research projects. Additionally, the position requires providing support for a moderate load of
industrial users. The position also provides opportunities to conduct independent research using the
microprobe as well as other facilities available at UMN.
An M.S. or Ph.D. degree in Earth Science or related field is required. This non-tenure track
position is available as a Research Fellow (M.S. degree, requisition #172657), or a Postdoctoral
Associate (up to 3 years experience post PhD, requisition #172658), or a Research Associate (3 years
post PhD experience, requisition #175602). To apply, candidates should go to
https://employment.umn.edu and attach a cover letter, resume (CV), statement of research interests,
and names/contact information of three references. Application review will begin on March 15,
2012, and will continue until the position is filled. For further information, contact Marc
Hirschmann, mmh-at-umn.edu.

Login Host: 58.169.36.89
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9, 25 -- From microscopylistserver-noreply-at-microscopy.com Wed Jan 11 21:00:31 2012
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Dear Melissa,

sectioning a polymer is not really a big deal if you have a (cryo)-
microtome. However, as always the devil is in the details!
First you need to know the TG (Glasstransition Temperature) of the
polymer in question. If it�s below RT you should prepare to do some
cryo cutting. If not, so much the better.
The real problem will be the ND, they will surely damage your knife
and will be bunked out of the polymer matirix to some extent.
However, you can try using a glass knife or I would suggest to use an
used diamond knife, where you don�t mind a few more additional
scratches.
Just give it a try, it�s not that difficult if you know how to
operate a microtome.

Cheers
Ingo
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} Email: mchimento-at-uab.edu Name: Melissa Chimento
}
} Organization: University of Alabama at Birmingham
}
} Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?
}
} Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
} introduce the NDs before polymerization at room temperature. They said that it forms a brick of
} polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
} NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
} suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
} the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
} they could check dispersion of the ND in the polymer with it??
}
}
} Thanks!
}
}
} Melissa Chimento
}
} University of Alabama at Birmingham-HRIF
}
} Department of Vision Science
}
} 205-934-1926
}
}
} Login Host: 138.26.156.111
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}
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Ingo Lieberwirth
Max Planck Institute for Polymer Research
Ackermannweg 10
D-55128 Mainz

Tel.: ++49 6131 379 580
Fax.: ++49 6131 379 100
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Colleagues,

Apologies if you receive this more than once as it is going to multiple
lists.

Over the last ~18 months, I have been setting up a new electron
microprobe lab at a 2800-student primarily undergraduate university in
West Virginia. Being curious if there might be any other undergraduate
schools with a fully-functioning probe lab, I prepared a compilation of
North American probe labs. Through this exercise, I think I have found
one other undergraduate probe lab.

Wanting to share, I have posted the compilation on our lab website at:
http://academics.concord.edu/microanalysis/OtherLabs.html

The compilation includes a Google Maps view as well as tables listing
the labs by institution name and by instrument type. Both academic and
government/industry labs are included. Also included are links to the
lab web sites where I have them.

Perhaps some of you may find this recent compilation to be useful.

I also could use your help in checking the accuracy and completeness of
the compilation. Please take a look and suggest additions/changes as
needed. I am sure that I have missed a few.

Thanks and best regards,

- Steve Kuehn

--

----------------
Dr. Stephen C. Kuehn
Research Assistant Professor
Manager, Electron Microprobe Facility & Tephra Lab
Science building, Room 106

Concord University
1000 Vermillion St
PO Box 1000, Campus Box F20
Athens, WV 24712-1000

http://academics.concord.edu/sckuehn/
http://academics.concord.edu/microanalysis/
sckuehn-at-concord.edu kuehnsc-at-gmail.com
304-384-6322

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Dear Melissa,
I agree with Ingo, my first try would cryo-UM depending on the polymer matrix. The only problem is
that the diamond particles are going to fall out of the material and leave nice big gouges in the
sections. Another possible solution would be a cryo-fracture and hi-res SEM of the material. You may
have to thin the brick down to a manageable thickness in order to fracture it. Good luck,
Vicky

Victoria M. Bryg
Research Associate - NCSER/USRA
NASA Glenn Research Center
216-433-9628

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Sent: Wednesday, January 11, 2012 10:20 PM
To: Bryg, Victoria M. (GRC-REC0)[National Center for Space Exploration Research]

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Email: mchimento-at-uab.edu Name: Melissa Chimento

Organization: University of Alabama at Birmingham

Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?

Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
introduce the NDs before polymerization at room temperature. They said that it forms a brick of
polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
they could check dispersion of the ND in the polymer with it??

Thanks!

Melissa Chimento

University of Alabama at Birmingham-HRIF

Department of Vision Science

205-934-1926

Login Host: 138.26.156.111
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Recently, we have noticed more reports of using Durcupan ACM resin in the literature, so we ordered a Durcupan ACM resin kit to try out. I noticed this resin is very viscous, much more so than the Embed-812 resin that we typically use. Does anyone have any particular resin infiltration schedules that have worked well for them for cell cultures and plant and animal tissues? I read that Durcupan causes less shrinkage than other resins, but does it have any other advantages over other resins for certain applications?

Thanks,
Shannon
Bio-Imaging Center
Delaware Biotechnology Institute
Newark, DE

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Hi Melissa

I would try SEM on a cry-fractured sample.

To fracture the sample, deep it in liquid nitrogen, until le nitrogen
stops to boil. Then, take it out and put in on an amboss, and hit it
with a hammer. To avoid the flakes to fly everywhere, you can put some
weeping paper on the amboss that you fold to cover the sample before you
hit it. You must do all these steps fast, to limit the reheating of the
sample.
It's not a very academic methode, nor it is very reproductible, but it
works. You can help the fracturing at a wanted place by making a notch
with a rasor blade. The way it will break depends of the type of the
polymer.

You choose in the flakes some flat pieces for the observation and coat
them with carbon. If the ND are really nano, you will need low energy
and high resolution. If the "nano" is in the 100 nm range or more, BSE
detector, higher energy (and maybe ESEM mode) should work.

Hope it helps

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Mat�riaux de Strasbourg
D�partement Surfaces et Interfaces
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail : Jacques.Faerber-at-ipcms.unistra.fr

Le 12/01/2012 04:22, microscopylistserver-noreply-at-microscopy.com a �crit :
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} Email: mchimento-at-uab.edu Name: Melissa Chimento
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} Organization: University of Alabama at Birmingham
}
} Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?
}
} Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
} introduce the NDs before polymerization at room temperature. They said that it forms a brick of
} polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
} NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
} suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
} the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
} they could check dispersion of the ND in the polymer with it??
}
}
} Thanks!
}
}
} Melissa Chimento
}
} University of Alabama at Birmingham-HRIF
}
} Department of Vision Science
}
} 205-934-1926
}
}
} Login Host: 138.26.156.111
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Perhaps instead of EM you might be better off investigating via
scattering, such as SAXS? That way you could look at the bulk sample and
see an average particle spacing, if the technique would permit it
(depends on density/spacing of NDs in the polymer).

Alternatively, instead of positive evidence (seeing the NDs) is your
work flexible enough to allow for the use of negative evidence instead
of positive evidence? In other words, if the NDs all get popped out
during microtoming you end up with a system full of voids. Investigation
of void spacing could indirectly give you info on ND spacing. This would
be dependent on how cleanly the NDs come out of the polymer - if they
tear or gouge the sample badly, then this would be a failed method.

-John Papalia

**********************
John M. Papalia, Ph.D.
Materials Scientist & Job seeker
**********************

On 1/12/2012 9:35 AM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Dear Melissa,
} I agree with Ingo, my first try would cryo-UM depending on the polymer matrix. The only problem is
} that the diamond particles are going to fall out of the material and leave nice big gouges in the
} sections. Another possible solution would be a cryo-fracture and hi-res SEM of the material. You may
} have to thin the brick down to a manageable thickness in order to fracture it. Good luck,
} Vicky
}
} Victoria M. Bryg
} Research Associate - NCSER/USRA
} NASA Glenn Research Center
} 216-433-9628
}
}
}
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} Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?
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} Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
} introduce the NDs before polymerization at room temperature. They said that it forms a brick of
} polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
} NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
} suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
} the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
} they could check dispersion of the ND in the polymer with it??
}
}
} Thanks!
}
}
} Melissa Chimento
}
} University of Alabama at Birmingham-HRIF
}
} Department of Vision Science
}
} 205-934-1926
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Use a FIB.

John Mardinly,
ASU

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Sent: Wednesday, January 11, 2012 8:22 PM
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Email: mchimento-at-uab.edu Name: Melissa Chimento

Organization: University of Alabama at Birmingham

Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?

Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
introduce the NDs before polymerization at room temperature. They said that it forms a �"brick�" of
polymer. I�'m not sure what to do with the sample. I�'ve had people bring in electro spun fibers with
NDs but not a solid chunk. From what I�'ve read polymers are extremely difficult to thin section. Any
suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEM�'s in
the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. I�'m not sure if
they could check dispersion of the ND in the polymer with it??

Thanks!

Melissa Chimento

University of Alabama at Birmingham-HRIF

Department of Vision Science

205-934-1926

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Melissa,

What size are the nanodiamonds are expected to be? If they in the 2 to 5 nm range, and the loading
fraction is not too high (~10% or lower by volume), microtoming should work just fine (cryo if Tg { RT).
At a 70 or 100 nm slice thickness, the vast majority should be well encapsulated with polymer, and
not drop out. A diamond microtome knife shouldn't get hurt either, unless the NDs are a bit bigger,
i.e., } 10 nm.
We do this routinely for some naturally-occurring polymer-nanodiamond samples, and haven't had to
change the diamond knife at all.

For } 10 nm NDs, I would go a with the FIB, as John said, either for direct-slice and view in the
FIB if you can get enough contrast, or for TEM imaging if contrast is a problem.
For the smaller NDs the surface damage and re-dep may make it hard to get the contrast you need for
either FIB-based or TEM-based imaging, unless you have a really good FIB and FIB operator.

Good luck.

Rhonda

On 1/11/2012 10:13 PM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?
}
} Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
} introduce the NDs before polymerization at room temperature. They said that it forms a brick of
} polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
} NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
} suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
} the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
} they could check dispersion of the ND in the polymer with it??
}
}
} Thanks!
}
}
} Melissa Chimento
}
} University of Alabama at Birmingham-HRIF
}
} Department of Vision Science
}
} 205-934-1926
}
}
} Login Host: 138.26.156.111
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--
__________________________________
Head, Nanoscale Materials Section
Code 6366
Naval Research Laboratory
4555 Overlook Avenue SW
Washington, DC 20375
(v) 202-404-4143
(f) 202-767-1697
rhonda.stroud-at-nrl.navy.mil
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Hi Melissa,

As a reality-sanity check, before preparing the polymer/diamond sample, I would suggest trying to
image a suspension of the nano-diamonds on a carbon film support. This is just to be certain of the
size of the nano-diamonds. Their size will influence how sliceable they may be with a diamond
knife. If they are very small, really nano sized, they may stay put in the polymer and make a nice
sample. The larger they are the higher the probability of pull out and knife damage. It would also
be good to ask what the researchers think the concentration of nano-diamond is in the polymer.

Years ago I had many researchers ask me to image what they thought were 5-10 nm diameter (lab-made)
nano-particles, these were often 100-700 nm diameter particles. This does not happen much anymore
but it is sometimes good to check before investing hours of your time. Also nano diamonds in
polymer may be easier to image with STEM dark field imaging rather than TEM--if you have the option.

They may want to try cryo-FIB of a block of their sample. It might be possible to get that done by
JEOL, FEI --not sure if Hitachi has a cryo FIB. IF the manufacturer can show off the results for
marketing purposes they will sometimes do exploratory samples for free.

Good luck,
Roseann

Roseann Csencsits, PhD
Scientist in Charge, Donner TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

On Jan 11, 2012, at 7:21 PM, microscopylistserver-noreply-at-microscopy.com wrote:

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} Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?
}
} Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
} introduce the NDs before polymerization at room temperature. They said that it forms a brick of
} polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
} NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
} suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
} the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
} they could check dispersion of the ND in the polymer with it??
}
}
} Thanks!
}
}
} Melissa Chimento
}
} University of Alabama at Birmingham-HRIF
}
} Department of Vision Science
}
} 205-934-1926
}
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I greatly appreciate all of the feedback I received to this post.
Thank you,

Melissa F. Chimento
University of Alabama at Birmingham
HRIF SHEL Room 912
1825 University Boulevard
Birmingham, AL 35294

LAB #: (205) 934-1926
OFFICE #: (205) 996-6469

mchimento-at-uab.edu

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Sent: Wednesday, January 11, 2012 9:20 PM
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Organization: University of Alabama at Birmingham

Title-Subject: [Filtered] Nano diamonds dispersed in a polymer TEM?SEM?

Message: We have a group that wants to see how nano diamonds(NDs) disperse in a polymer. They
introduce the NDs before polymerization at room temperature. They said that it forms a brick of
polymer. Im not sure what to do with the sample. Ive had people bring in electro spun fibers with
NDs but not a solid chunk. From what Ive read polymers are extremely difficult to thin section. Any
suggestions about how to evaluate ND dispersion in a solid would be appreciated. We have 2 TEMs in
the lab. Should I try cryo-sectioning the material? We have a new ESEM on campus. Im not sure if
they could check dispersion of the ND in the polymer with it??

Thanks!

Melissa Chimento

University of Alabama at Birmingham-HRIF

Department of Vision Science

205-934-1926

Login Host: 138.26.156.111
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Any advice for EBSD of zircaloys? I tried multi-hour colloidal silica (no patterns, 15-30 kV) and then tried a few minutes of colloidal silica+ammonium hydroxide + hydrogen peroxide (still no patterns, 15-30 kV).

Has anyone had good luck? My next resort is our Gatan Ilion ion polisher, but I'm hoping for a simple answer to let me use my already mounted and polished specimens.

Thanks,
Chad Parish

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Dear All,
maybe somebody out there can help me with classifying this diatomee:
http://www.electronmicroscopy.info/diatomee/index.htm

I found it on a ca 300 year old piece of slag from iron smelting in a valley in the northern part of Bavaria, Germany. The small
river I found it in is not too warm during the year, height above MSL is ca. 800 to 400 meter.
Google earth link is at
http://www.electronmicroscopy.info/diatomee/iron_slug.kmz

Thanks,
Stefan

--

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Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
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++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
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www.assisi.de
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Anfahrt: http://Mail.map24.com/Stefan.Diller
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That a great way to start a work week: A diatom question! Unfortunately I don't have any of my diatom references at work, but I have a question of my own.

I cutting cryo-thin sections of silica loaded silicone rubber with a diamond histology knife (it's all I have). It as a Tg of -127C. I'm working dry at -140C using sugar water to pick-up the thin sections for TEM imaging at 80kv. I don't know the loading of silica in the silicone polymer. The microtome is a RMC microtome with cryo-stage.

It's not going well. I can't seem to get to 100 nm. The color guide, based on the interference of light, suggests I at 280nm. In the few thin spots or tears I can see the particles of silica I'm after but the rest of the section is too thick.

Has anyone practical experience with cryo microtoming of silica loaded silicone?
I'm open to ideas

Thanks....
Frank

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Stefan,
I pulled out my only reference to European diatoms, Heurch's Treatise on Diatomaceae published in 1896. I just have a photographic copy of it so the images aren't that sharp and the line drawings are so-so but I think your diatom best matches Navicula viridula.

hope this helps .....
Frank

-----Original Message-----
X-from: stefan.diller-at-t-online.de [mailto:stefan.diller-at-t-online.de]
Sent: Monday, January 16, 2012 7:00 AM
To: Frank Karl

Dear All,
maybe somebody out there can help me with classifying this diatomee:
http://www.electronmicroscopy.info/diatomee/index.htm

I found it on a ca 300 year old piece of slag from iron smelting in a valley in the northern part of Bavaria, Germany. The small
river I found it in is not too warm during the year, height above MSL is ca. 800 to 400 meter.
Google earth link is at
http://www.electronmicroscopy.info/diatomee/iron_slug.kmz

Thanks,
Stefan

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Dear Listers
Looking at a mucosal animal epithelium with SEM, I came across a creature
that is strange to me. It's like 30 microns long tube at the shape of L,
the short arm has a hole at the end and the long one is twisted like a
screw. Maybe it is a short of borrelia? Whoever is familiar with
microorganisms, please have a look at

http://www.eikonika.net/v2/photo_list_nikas.php

Any comments will be greatly appreciated!

Best regards
yorgos

Dr Yorgos Nikas
Athens Innovative Microscopy
Skra 36 Voula 16673 GREECE

Tel/fax +30 210 8957677
mobile +30 6945 107477
eikonika-at-otenet.gr

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Thanks all for your help concerning my diatomee problem.

The most probable outcome is "Hippodonta capitata", as suggested by Prof. Hans Rudolf Thierstein.
He also pointed me to a really impressing source on diatomee:
http://westerndiatoms.colorado.edu/taxa/species/hippodonta_capitata

Best wishes,
Stefan

--

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Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
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www.assisi.de
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Maybe this question would be best for Professor Thierstein, but is there a good online guide for classifying diatoms?

I have looked at quite a few diatoms in water samples over the years but have not embarked on trying to classify them. They were simply a significant part of the solids load of the water sample. Many looked like your sample.

However, from a quick search prompted by your question, I see similarities between hippodonta, planothidium, and navicula. The differences appear to be subtle between them. Therefore, I wonder about a good, definitive classification scheme.

Warren Straszheim

-----Original Message-----
X-from: stefan.diller-at-t-online.de [mailto:stefan.diller-at-t-online.de]
Sent: Tuesday, January 17, 2012 9:38 AM
To: wesaia-at-iastate.edu

Thanks all for your help concerning my diatomee problem.

The most probable outcome is "Hippodonta capitata", as suggested by Prof. Hans Rudolf Thierstein.
He also pointed me to a really impressing source on diatomee:
http://westerndiatoms.colorado.edu/taxa/species/hippodonta_capitata

Best wishes,
Stefan

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This, and many more Diatomacea tracts, might also help - even if specified for US.

http://books.google.com/ebooks?id=G35IAAAAYAAJ&dq=diatoms%20of%20united%20states%20subject%3A%22Science%22&lr&as_brr=5&ei=56oVT8eHDoq0NtiY3LsG&source=webstore_bookcard

http://books.google.com/ebooks?id=mzYDAAAAQAAJ&dq=diatoms%20of%20united%20states%20subject%3A%22Science%22&lr&as_brr=5&ei=26sVT4eAKIz6M5KvpOoB&source=webstore_bookcard Paleontology by Richard Owen

http://books.google.com/ebooks?id=rZY_AAAAYAAJ&dq=diatoms%20of%20united%20states%20subject%3A%22Science%22&lr&as_brr=5&ei=ickVT-WcJoPUM4e8re0L&source=webstore_bookcard (A Treatise on the Diatomacea)

Cheers,

FCM

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

If one looks long enough, one may actually see something frozen in time that flashes one's mind to a new state.
1. The day that I asked myself how the hydrogen atom's mass could actually be: 1.0008.
2. The day, while looking at a slide of some tissue, I saw a leukocyte that had been caught sneaking in or out of a capillary: http://www.annualreviews.org/doi/abs/10.1146/annurev-pathol-011110-130224

-----Original Message-----
X-from: stefan.diller-at-t-online.de [mailto:stefan.diller-at-t-online.de]
Sent: Tuesday, January 17, 2012 10:44 AM
To: Monson, Frederick

Thanks all for your help concerning my diatomee problem.

The most probable outcome is "Hippodonta capitata", as suggested by Prof. Hans Rudolf Thierstein.
He also pointed me to a really impressing source on diatomee:
http://westerndiatoms.colorado.edu/taxa/species/hippodonta_capitata

Best wishes,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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Title-Subject: [Filtered] Wanted, TEM

Message: Small laboratory in New Jersey is looking to purchase a well maintained used TEM.
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Hi all,
A student wants to use high pressure freezing on dinoflagellates (size
approx. 10 microns). Has anyone done this? If yes, did you wick the
dinos into cellulose capillary tubes prior to freezing? Or, is there a
better method for keeping track of them?

thanks,
Beth

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Having installed a FEG image analyzer and comparing its results with our tungsten SEM, we are seeing a slight bias with respect to brighter particles measuring smaller (possibly more accurately) with the FEG. The image is backscattered electrons and we use a fixed background threshold for isolating particles from the epoxy. It occurred to me that clipping the edge at the same background value would make brighter particles larger depending on how "fuzzy" the edge is. It would even be possible to predict this "bright versus dark" bias if the SEM's spot size was determined. Some difference should be associated with each SEM's magnification calibration, but I'm accounting for that by first relating calibration to the darkest minerals (quartz which is abundant).

I thought before I re-invent the wheel and get involved with the math and all the other possible variables (e.g., pixel resolution, irregular perimeters, ...) that I'd ask my peers if this has been done before(?) ... leastwise, I haven't yet found a reference ...

TIA & Cheerios from the Avalon
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
Information: http://www.mun.ca/creait/maf/
Scheduling: http://www.mun.ca/creait/maf/SEM-MLA/calendar.php
(709) 864-6799 (Ofc)
(709) 864-6790 (Lab)
cogito ergo ZzoooomM

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5, 25 -- From michael-at-shaffer.net Wed Jan 18 13:30:06 2012
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That doesn't quite make sense to me. I don't see how changing to a higher resolution would lead to what you describe.

In a previous research life, I did a lot of image analysis on minerals in coal. The brighter ones (primarily pyrite) were cut much lower than half max when the threshold was set to detect the darker minerals which comprised the bulk of the mineral matter. Therefore, I expected the pyrite to be measured larger than actual size. The amount of error related to the spot size and the "fuzziness" of the signal. However, the effect was far beyond what I would expect the difference to be between W and field-emission guns.

I would look into other things that might have changed with the changeover.

Is the response of the BSE signal the same, or have they introduced some non-linearity that might have changed how the threshold is set? I would be interested in seeing the relative brightness of the various minerals under similar setups.

Has the software adapted some sort of local thresholding? I know I would have appreciated that back in the day, but we didn't have the computing power to well implement it.

I'd be interested in continuing the conversation off-line if that would be more expedient.

Warren Straszheim
former operator of a LeMont Scientific DB-10 image analyzer, circa 1981.
________________________________________
X-from: michael-at-shaffer.net [michael-at-shaffer.net]
Sent: Wednesday, January 18, 2012 1:30 PM
To: wesaia-at-iastate.edu

Having installed a FEG image analyzer and comparing its results with our tungsten SEM, we are seeing a slight bias with respect to brighter particles measuring smaller (possibly more accurately) with the FEG. The image is backscattered electrons and we use a fixed background threshold for isolating particles from the epoxy. It occurred to me that clipping the edge at the same background value would make brighter particles larger depending on how "fuzzy" the edge is. It would even be possible to predict this "bright versus dark" bias if the SEM's spot size was determined. Some difference should be associated with each SEM's magnification calibration, but I'm accounting for that by first relating calibration to the darkest minerals (quartz which is abundant).

I thought before I re-invent the wheel and get involved with the math and all the other possible variables (e.g., pixel resolution, irregular perimeters, ...) that I'd ask my peers if this has been done before(?) ... leastwise, I haven't yet found a reference ...

TIA & Cheerios from the Avalon
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
Information: http://www.mun.ca/creait/maf/
Scheduling: http://www.mun.ca/creait/maf/SEM-MLA/calendar.php
(709) 864-6799 (Ofc)
(709) 864-6790 (Lab)
cogito ergo ZzoooomM

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==============================================================================================================
Good Morning,

Dear Amy,
just to let you know:
The whole LKB-product branch has been taken over some 10 years ago by LEICA (cf. http://www.leica-microsystems.com/).
So you'll find some results if you search "Google" by the phrase
} LEICA 2178 KnifeMaker II { or } LEICA 2178 KnifeMaker II AND Manual {.

Since I have not found the Type you were asking for in my own PC-files (Reichert, LKB, LEICA, ZEISS, etc.)
I googled in my own interest and therefore I - exceptionally and hopefully tolerated by the listers - communicate those results I found quickly (and perhaps could be a short way out of your dilemma):

Within the first 20 results there were no results for a {MANUAL available}

but, perhaps you could be supported by EM or Imaging Lab's posessing such a KnifeMaker Type:
cf: http://www.memphis.edu/imc/services.htm a Lab which is - unfortunately - some 500 miles away from your location:

Staff communication
http://www.memphis.edu/imc/staff.htm :
Dr. Omar Skalli
Director
oskalli-at-memphis.edu
Phone: (901) 678-2034
Office: 101 Life Science Bldg

Further information on use of the IMC, including fee schedules, can be obtained by contacting Ms. Lou G. Boykins, Laboratory Coordinator at (901) 678-4233 (phone); (901) 678-4457 (FAX) or lgboykns-at-memphis.edu
Ms. Lou Boykins
Laboratory Coordinator
Phone: (901) 678-2034
Office: 101 Life Science Bldg

Ms. Renada Scott
Research Technician II
rjscott3-at-memphis.edu
Phone: (901) 678-2034
Office: 101 Life Science Bldg
Perhaps they can send you a copy of their manual.

At the moment of writing this I do not know about a positive response from any Listserver member.

Another source would be perhaps:

Material Testing - Page 5 - Manuals - HiTechTrader.com
see: http://www.hitechtrader.com/dspManuals.cfm?1=1&catID=2262&MaxRows=25&sorder=ASC&sort=m.Make&page=5

They are listing:
"This instruction manual describes the function and operation of the LKB 2178 KnifeMaker II.
LKB 2178 KnifeMaker II Instruction Manual
Make: LKB
Model: 2178
Reference No: M-050110204
This instruction manual describes the function and operation of the LKB 2178 KnifeMaker II
BUT: Disclaimer for all manuals: Manuals listed below are not for copy, sale and/or redistribution. The manuals listed below are used by HiTechTrader.com to repair and maintain equipment. Please contact us (== cf: http://www.hitechtrader.com/contact.cfm?PgID=115 ) if you need any assistance "

Hoping this perhaps was/is of help to you,

best regards,
Wolfgang MUSS PhD
EM-Lab
Univ. Inst. Pathology
SALZBURG-AUSTRIA

=======================================

Von: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Gesendet: Donnerstag, 19. J�nner 2012 00:40
An: Mu� Wolfgang
Betreff: [Microscopy] manual for a LKB 2178 Knifemaker
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I was wondering if anyone knew where to find a manual for a LKB 2178 Knifemaker II?
So far, google hasnt been all that helpful.
Thanks!

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Email: acamacho-at-rd.us.loreal.com Name: Alejandra Camacho

Organization: Sr Research Scientist at L'Oreal

Title-Subject: [Filtered] Cathodoluminescence

Message: Hello dear colleagues,
I'm seeking your help because I have two mineral samples with same
overall composition (according to my EDS, XRD's are also the same for
both) but have different provenance and different performance. I believe
the difference is in some trace elements but I need to confirm. I would
like to try cathodoluminescence before any exotic technique like neutron
activation or PIXE but I don't have a CL detector in house, can you
recommend any lab or institution where I could get a couple of samples
analyzed? (I apologize for introducing a bit of a commercial here but I
haven't been very succesful in finding this info)
Thanks in advance for you advice

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Good Morning,
Let me recommend MicroTrace to you. They are north of Chicago and I believe they can help you out. Here's a link:

http://www.microtracescientific.com/

Good luck........
Frank Karl
ARDL

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Email: acamacho-at-rd.us.loreal.com Name: Alejandra Camacho

Organization: Sr Research Scientist at L'Oreal

Title-Subject: [Filtered] Cathodoluminescence

Message: Hello dear colleagues,
I'm seeking your help because I have two mineral samples with same
overall composition (according to my EDS, XRD's are also the same for
both) but have different provenance and different performance. I believe
the difference is in some trace elements but I need to confirm. I would
like to try cathodoluminescence before any exotic technique like neutron
activation or PIXE but I don't have a CL detector in house, can you
recommend any lab or institution where I could get a couple of samples
analyzed? (I apologize for introducing a bit of a commercial here but I
haven't been very succesful in finding this info)
Thanks in advance for you advice

Login Host: 198.16.3.248
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Barbara,

I had the same problem making a 2% solution about 20 years ago - using old UA. As soon as I started using a new supply the UA went into solution as I thought it should.

If your UA comes in a can it is not a bad idea to store the bottle in the can between uses. I have found that this extends the useable life of the crystals until I empty the bottle.

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E - Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}

Opinions and experiences related are those of Pat Connelly and do not represent the NIH. This message is not confidential and can be freely shared and reproduced.

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Message: I have been having a problem with getting uranyl acetate to go
into solution. I have been using the same bottle of UA for over 22
years. I have not had any problems when I was using nanopure water, but
now that I am using distilled water, I can't seem to make a 3% solution
of UAaq. Is there anything I could do to help it go into solution? I am
near the end of the bottle. Maybe I should just buy another bottle? Ot
try to get the lab to get the nanopure water working again ( I think
they need a special filter). Or would adding a little methanol help? I
am open to suggestions! Thank you.

Login Host: 138.9.63.239
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==============================Original Headers==============================
11, 30 -- From connellyps-at-nhlbi.nih.gov Fri Jan 20 09:00:24 2012
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Hi,
I have no experiencing with aging uranyl acetate. But am
curious how time could cause a crystal to become insoluble. Many
chemicals on the lab shelf absorb water over time (they often form
cakes). This make them difficult to weigh out but I cannot think why
it would hinder solubility. Alternatively, could an oxide be forming
on the surface of the grains, an oxide might in insoluble (think
rust). If the UA powder is not super fine, I guess you could try to
grind it with a mortar and pestle to expose fresh surface, although
from a safety point of you that seems dubious (UA aerosol -- yumm!).
Anyway, I'd be interested to hear if anyone actually knows how time
leads to insolubility (of powders -- not mentality, smile).

As ever
Tobias

At 9:01 AM -0600 1/20/12, connellyps-at-nhlbi.nih.gov wrote:
}
}
} Barbara,
}
} I had the same problem making a 2% solution about 20 years ago -
} using old UA. As soon as I started using a new supply the UA went
} into solution as I thought it should.
}
} If your UA comes in a can it is not a bad idea to store the bottle
} in the can between uses. I have found that this extends the useable
} life of the crystals until I empty the bottle.
}
} Pat
}
} Patricia Stranen Connelly
} Research Assistant
} NHLBI Electron Microscopy Core
} National Institutes of Health
} 14 Service Road West
} Bldg. 14E - Rm. 111B MSC 5570
} Bethesda, MD 20892-5570
} Phone 301-496-3491
} FAX 301-402-0170
} connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}
}
} Opinions and experiences related are those of Pat Connelly and do
} not represent the NIH. This message is not confidential and can be
} freely shared and reproduced.
}
}
} ---------------------------------------------------------------------------
} Email: bplowman-at-pacific.edu Name: Barbara Plowman
} Organization: University of the Pacific, Arthur A. Dugoni School of
} Dentistry
} Title-Subject: [Filtered] Uranyl acetate
}
} Message: I have been having a problem with getting uranyl acetate to go
} into solution. I have been using the same bottle of UA for over 22
} years. I have not had any problems when I was using nanopure water, but
} now that I am using distilled water, I can't seem to make a 3% solution
} of UAaq. Is there anything I could do to help it go into solution? I am
} near the end of the bottle. Maybe I should just buy another bottle? Ot
} try to get the lab to get the nanopure water working again ( I think
} they need a special filter). Or would adding a little methanol help? I
} am open to suggestions! Thank you.
}
} Login Host: 138.9.63.239
} =============================

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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5, 21 -- From: Tobias Baskin {baskin-at-bio.umass.edu}
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Dear Barbara,
I think this problem has to do with the pH of the water that is used. Probably the pH of the nanopure water was higher than the distilled water.

Groeten, Hans Janssen.

-----Original Message-----
X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
Sent: vrijdag 20 januari 2012 16:30
To: Hans Janssen

Hi,
I have no experiencing with aging uranyl acetate. But am
curious how time could cause a crystal to become insoluble. Many
chemicals on the lab shelf absorb water over time (they often form
cakes). This make them difficult to weigh out but I cannot think why
it would hinder solubility. Alternatively, could an oxide be forming
on the surface of the grains, an oxide might in insoluble (think
rust). If the UA powder is not super fine, I guess you could try to
grind it with a mortar and pestle to expose fresh surface, although
from a safety point of you that seems dubious (UA aerosol -- yumm!).
Anyway, I'd be interested to hear if anyone actually knows how time
leads to insolubility (of powders -- not mentality, smile).

As ever
Tobias

At 9:01 AM -0600 1/20/12, connellyps-at-nhlbi.nih.gov wrote:
}
}
} Barbara,
}
} I had the same problem making a 2% solution about 20 years ago -
} using old UA. As soon as I started using a new supply the UA went
} into solution as I thought it should.
}
} If your UA comes in a can it is not a bad idea to store the bottle
} in the can between uses. I have found that this extends the useable
} life of the crystals until I empty the bottle.
}
} Pat
}
} Patricia Stranen Connelly
} Research Assistant
} NHLBI Electron Microscopy Core
} National Institutes of Health
} 14 Service Road West
} Bldg. 14E - Rm. 111B MSC 5570
} Bethesda, MD 20892-5570
} Phone 301-496-3491
} FAX 301-402-0170
} connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}
}
} Opinions and experiences related are those of Pat Connelly and do
} not represent the NIH. This message is not confidential and can be
} freely shared and reproduced.
}
}
} ---------------------------------------------------------------------------
} Email: bplowman-at-pacific.edu Name: Barbara Plowman
} Organization: University of the Pacific, Arthur A. Dugoni School of
} Dentistry
} Title-Subject: [Filtered] Uranyl acetate
}
} Message: I have been having a problem with getting uranyl acetate to go
} into solution. I have been using the same bottle of UA for over 22
} years. I have not had any problems when I was using nanopure water, but
} now that I am using distilled water, I can't seem to make a 3% solution
} of UAaq. Is there anything I could do to help it go into solution? I am
} near the end of the bottle. Maybe I should just buy another bottle? Ot
} try to get the lab to get the nanopure water working again ( I think
} they need a special filter). Or would adding a little methanol help? I
} am open to suggestions! Thank you.
}
} Login Host: 138.9.63.239
} =============================

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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13, 25 -- From j.janssen-at-nki.nl Fri Jan 20 09:48:40 2012
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I know that uranyl acetate is generally considered as only slowly/poorly soluble in water. I would suspect that acetate salts would decompose more readily than some other salts and so I had always just assumed that old uranyl acetate just slowly oxidised producing a less soluble mixture.

Certainly I had heard that 10+ year old uranyl acetate was less soluble than a fresh supply so I had always refused kind gifts of uranyl acetate if it was old.

I don't know if this helps much.

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: j.janssen-at-nki.nl [j.janssen-at-nki.nl]
Sent: 20 January 2012 15:56
To: Malcolm Haswell

Dear Barbara,
I think this problem has to do with the pH of the water that is used. Probably the pH of the nanopure water was higher than the distilled water.

Groeten, Hans Janssen.

-----Original Message-----
X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
Sent: vrijdag 20 januari 2012 16:30
To: Hans Janssen

Hi,
I have no experiencing with aging uranyl acetate. But am
curious how time could cause a crystal to become insoluble. Many
chemicals on the lab shelf absorb water over time (they often form
cakes). This make them difficult to weigh out but I cannot think why
it would hinder solubility. Alternatively, could an oxide be forming
on the surface of the grains, an oxide might in insoluble (think
rust). If the UA powder is not super fine, I guess you could try to
grind it with a mortar and pestle to expose fresh surface, although
from a safety point of you that seems dubious (UA aerosol -- yumm!).
Anyway, I'd be interested to hear if anyone actually knows how time
leads to insolubility (of powders -- not mentality, smile).

As ever
Tobias

At 9:01 AM -0600 1/20/12, connellyps-at-nhlbi.nih.gov wrote:
}
}
} Barbara,
}
} I had the same problem making a 2% solution about 20 years ago -
} using old UA. As soon as I started using a new supply the UA went
} into solution as I thought it should.
}
} If your UA comes in a can it is not a bad idea to store the bottle
} in the can between uses. I have found that this extends the useable
} life of the crystals until I empty the bottle.
}
} Pat
}
} Patricia Stranen Connelly
} Research Assistant
} NHLBI Electron Microscopy Core
} National Institutes of Health
} 14 Service Road West
} Bldg. 14E - Rm. 111B MSC 5570
} Bethesda, MD 20892-5570
} Phone 301-496-3491
} FAX 301-402-0170
} connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}
}
} Opinions and experiences related are those of Pat Connelly and do
} not represent the NIH. This message is not confidential and can be
} freely shared and reproduced.
}
}
} ---------------------------------------------------------------------------
} Email: bplowman-at-pacific.edu Name: Barbara Plowman
} Organization: University of the Pacific, Arthur A. Dugoni School of
} Dentistry
} Title-Subject: [Filtered] Uranyl acetate
}
} Message: I have been having a problem with getting uranyl acetate to go
} into solution. I have been using the same bottle of UA for over 22
} years. I have not had any problems when I was using nanopure water, but
} now that I am using distilled water, I can't seem to make a 3% solution
} of UAaq. Is there anything I could do to help it go into solution? I am
} near the end of the bottle. Maybe I should just buy another bottle? Ot
} try to get the lab to get the nanopure water working again ( I think
} they need a special filter). Or would adding a little methanol help? I
} am open to suggestions! Thank you.
}
} Login Host: 138.9.63.239
} =============================

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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University of Sunderland - life-changing: see our new TV advert at
http://www.lifechangingsunderland.com or http://www.sunderland.ac.uk

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Our color printer, which I use to print B-W copies of TEM images of carbon black, is sounding like a rusty gate. On the chance we
replace it, I'm looking for suggestions: What's going to give me the nicest B+W TEM print? Any suggestions? Money is (and when is it not?) a factor.

If you're shy, contact me directly, or not. All suggestions will be considered,

Thanks...................

Frank

Frank Karl
Microscopist
ARDL
2887 Gilchrist Road
Akron, Ohio 44305
330-794-6600

________________________________
This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Reply to Hans.

I had used the same water source with both samples of UA , old and new. I know that the pH stayed the same most of the year so I do not think that the pH changed from morning to afternoon of the same day. If I remember correctly our pH usually ran in the 6.5 range. I know it was well below a pH of 7.0 any time I tested it.

Reply to Malcolm.

I had trouble getting the old UA to go into a 2% solution over a period of time. It just kept getting worse to the point that I had it mixing for up to two days (under a metal can to exclude the room lights) which had me questioning what was wrong when I knew that others made 4% UA with no difficulty. At that time I "borrowed" a gram from another lab which I knew was going into solution fine according to my EM Tech. friend. I then used my water, glass bottle, etc. the same that I had done before with my old UA. It went into solution within a short period of time so I ordered a new supply of UA from the same supplier of my old bottle and the borrowed UA and have not had a problem since.

I suggest that those who offer me gifts of old UA send the bottles out the next time they have a pick up of radioactive waste. I have tried several old supplies over the years and found none to go into solution readily.

Reply to Tobias.

When I started in EM back in 1971 my supply of UA was of coarse crystals which I was instructed to grind fine. I had a dedicated mortar and pestle for this purpose. I was very glad that when I ordered my first new bottle of UA some years later that it came as a fine, almost powder like grade of crystals. This is how my UA has come since that time.

Since it is now 2012 and I am still alive I guess I did a good job of not inhaling the dust! According to a very old edition of the Merc Index, UA was used as a snuff. Hard to believe with all the cautions on current MSDS sheets.

Pat Connelly

Opinions and experiences related are those of Pat Connelly and do
not represent the NIH. This message is not confidential and can be
freely shared and reproduced.

________________________________
X-from: {j.janssen-at-nki.nl}
Reply-To: {j.janssen-at-nki.nl}

Well, this has been an interesting discussion about what happens with
aging uranyl acetate (UAc) salts.

Although I am certainly not a chemist, I suspect several events (some
already mentioned by other contributors) are taking place to cause the
salts to become less soluble:

1. Degradation of acetate. Acetates are notoriously unstable and
break down with time. If you detect a strong smell of acetic acid
(carefully, by wafting the air above the container), then
decomposition is taking place.

2. Phytolytic decomposition. Most people keep the UAc away from the
light; however, this is not always the case. I've seen solutions
sitting on top of counters for long periods of time in clear, glass
containers. You should cover the container with aluminum foil and keep
it refrigerated (to cut down on light even more).

3. Radiolytic decomposition. UAc (even depleted, U238 Ac) is weakly
radioactive, giving off alpha, but also beta particles and gamma rays
as part of the decay process. Over time, these will degrade most
chemical compounds. In fact, ever wonder what was coating the inside
of your UAc staining vessel? That insoluble material is uranyl oxide
(JIÍ TEPLÝ & VÁCLAV TULÍK. 1963. Formation of Peroxidic Precipitate in
the Radiolysis of Uranyl Nitrate Ketone Solutions. Nature 200:
671-672).

Now, for some "popular" trivia. Anyone ever hear of a Revigator?

That's a water vessel lined with uranium that releases radon into the
water! Back in the 1920's they were sold to fitness folks to "restore
water's lost element" and invigorate the body……… Frightening! You can
occasionally find them on EBay and I am surprised they are even
allowed to be sold.

--
John J. Bozzola, Ph.D., (Happily Retired) Professor & Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
750 Communications Drive
Carbondale, IL 62901
Phone: 618-453-3730

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Hi Listers,

It looks like I am becoming illegally blind: I cannot find instructions for authors and/or abstract template on M&M site. Recommended in Call for Papers link does not connect to the right page.

Were there any changes in template or I can use my old abstract as a template?

Thanks,

Vladimir

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Note: I have already sent the O.P. Charles Humphrey's Alcian Blue
method (which he notes was published by Nermut in 1982).

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} Date: Mon, 23 Jan 2012 05:54:13 -0800
} From: {AssociationManagement-at-microscopy.org}
} Reply-To: Amanda {lever.amanda1-at-gmail.com}
} Subject: Ask-A-Microscopist
}
} Below is the result of your form, submitted on Monday, January 23,
} 2012 at 05:53:59 AM.
}
} realname - Amanda
} Email - lever.amanda1-at-gmail.com
} ORGANIZATION - Draper Laboratory
} EDUCATION - Graduate College
} LOCATION - Boston, MA
} SUBJECT_OF_QUESTION - Transmission Electron Microscopy
} QUESTION - I am currently using a UA negative stain on carbon grids
} to visualize and image liposomes. I have been noticing heterogeniety
} across the EM grids (some liposomes look nice and maintain the 3D
} shape while others appear flat and 2D in shape). Why might this be
} happening and are there any techniques to prevent it ?
}

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Note: I have referred the O.P. to MSA's surplus equipment page, but
anything specific to what he's looking for would help him.

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} realname - Ralph Appy
} Email - r.appy-at-cox.net
} ORGANIZATION - Cabrillo Marine Aquarium
} LOCATION - San Pedro, CA, U.S.A.
} SUBJECT_OF_QUESTION - New/Refurbished Microscope
} QUESTION - I am a retired environmental scientist and rekindling my
} interest in fish parasitology at a local marine aquarium. I would
} like to find a used/refurbished phase contrast microscope with a
} drawing tube. Not sure if any members might have such a surplus
} microscope. Not having much luck on what I want on the internet. I
} know this is an unusual request. I am bonafide [...just published
} in J Parasitol 97(6):1035-1048] and not indepentently wealthy.
}
} Ralph

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Greetings Fellow Microscopists,

The Hooke College of Applied Sciences, located in Westmont, IL, is offering its transmission electron microscopy short course March 6-8, 2012. In addition to lectures, this course emphasizes hands-on training using state-of-the-art equipment. For further details and registration information, please follow the link below:

http://www.hookecollege.com/courses/course.asp?COURSE_ID=53

Best regards,

Elaine Schumacher

*********************************************************************
Elaine F.Schumacher
Senior Research Scientist
McCrone Associates, Inc.
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7417 (fax)
E-mail: eschumacher-at-mccrone.com
Web Site: www.mccrone.com

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This message and any attachments are solely for the
intended recipient. If you are not the intended recipient,
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I have a sample of zinc borate that I've been asked to determine the atomic number ratio of boron to zinc. Has anybody had any luck in detecting boron and if so, how did you go about it? I have IXRF software using an Amray 1830 with a light element detector. Is it a matter of setting up the detector threshold so you can see below carbon or what? Any help would be greatly appreciated!

Don Kierstead
Microscopist
ARDL, Inc.
www.ardl.com
(866) 778-ARDL [toll free]
(330) 794-6600 [worldwide]
(330) 794-6610 [fax]

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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5, 25 -- Subject: RE: Detection of Boron by EDX
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The first thing you do is proceed with great caution. If your system can detect boron, I am sure it will spit out a number. It might even have some resemblance to the truth, but probably not.

The second thing you should probably do is start looking into the difficulty of light element analysis. You have boron and oxygen (light elements) combined with zinc (a rather heavy element). Look into the ZAF factors and see how big the absorption corrections. Ask yourself how well you know the mass absorption coefficients for those combinations. See what you can do about changing conditions (dropping voltage) to reduce the magnitude of the corrections.

The third thing you do would be to make sure your system can see the boron peak. My guess is that you are probably not optimized to detect boron. We had an old Kevex Quantum detector running on one of the early IXRF systems. You might be able to adjust your discriminators to better see the peak. We had to. In the end, we could see most of the boron peak, but not much if any of the background on the low energy side.

The fourth thing would be to get some reference materials. If your system cannot give you right answers on a known compound (and it probably won't), then you can forget about trying to analyze an unknown compound. You may be able to collect your own standards to help the matter, but that is not a job for novices. (If you succeed, you may have progressed beyond the novice stage. There is much more to be said on this topic.)

Another thing is to consider the limitations of EDS in general. I suppose they will not have a flat, polished sample. You'll need to consider the effects of geometry. They affect results and become more significant the lighter the elements. Depending on where you hit a particle (on the side near the detector or the side away), you can probably produce any results you want.

Finally, I would get a large supply of salt so you can hand out your results with many grains. Seriously, I would try to get your client to accept qualitative results such as "The sample shows Zn, B, and O. It could be borate." This will be even more convincing if you can compare spectra of their unknown to a known sample of zinc borate and say, "See, they match". That may be as far as you really want to push it.

Regards,
Warren Straszheim

-----Original Message-----
X-from: donk-at-ardl.com [mailto:donk-at-ardl.com]
Sent: Monday, January 23, 2012 10:42 AM
To: wesaia-at-iastate.edu

I have a sample of zinc borate that I've been asked to determine the atomic number ratio of boron to zinc. Has anybody had any luck in detecting boron and if so, how did you go about it? I have IXRF software using an Amray 1830 with a light element detector. Is it a matter of setting up the detector threshold so you can see below carbon or what? Any help would be greatly appreciated!

Don Kierstead
Microscopist
ARDL, Inc.
www.ardl.com
(866) 778-ARDL [toll free]
(330) 794-6600 [worldwide]
(330) 794-6610 [fax]

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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I believe I would first consider surface tension as the culprit. Are all liposomes identically resistant to the passage of the air-water interface?

Hope this at least serves as a last resort,

Fred Monson

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

If one looks long enough, one may actually see something frozen in time that flashes one's mind to a new state.
1. The day that I asked myself how the hydrogen atom's mass could actually be: 1.0008.
2. The day, while looking at a slide of some tissue, I saw a leukocyte that had been caught sneaking in or out of a capillary: http://www.annualreviews.org/doi/abs/10.1146/annurev-pathol-011110-130224

-----Original Message-----
X-from: oshel1pe-at-cmich.edu [mailto:oshel1pe-at-cmich.edu]
Sent: Monday, January 23, 2012 9:18 AM
To: Monson, Frederick

Note: I have already sent the O.P. Charles Humphrey's Alcian Blue method (which he notes was published by Nermut in 1982).

***************************************************************************************
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} Date: Mon, 23 Jan 2012 05:54:13 -0800
} From: {AssociationManagement-at-microscopy.org}
} Reply-To: Amanda {lever.amanda1-at-gmail.com}
} Subject: Ask-A-Microscopist
}
} Below is the result of your form, submitted on Monday, January 23,
} 2012 at 05:53:59 AM.
}
} realname - Amanda
} Email - lever.amanda1-at-gmail.com
} ORGANIZATION - Draper Laboratory
} EDUCATION - Graduate College
} LOCATION - Boston, MA
} SUBJECT_OF_QUESTION - Transmission Electron Microscopy QUESTION - I am
} currently using a UA negative stain on carbon grids to visualize and
} image liposomes. I have been noticing heterogeniety across the EM grids
} (some liposomes look nice and maintain the 3D shape while others appear
} flat and 2D in shape). Why might this be happening and are there any
} techniques to prevent it ?
}

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Hi All,

I've found a couple of papers that list Pt-blue as an alternative to Uranyl
Acetate, but I haven't been able to track down the way to make it. Does
anyone have a working recipe as how to make the Pt-blue? Thanks for any
help/tips.

--
Bryan Hansen
EM Technician
Rocky Mountain Laboratory
NIAID/NIH
bryan.todd.hansen-at-gmail.com
hansenbry-at-niaid.nih.gov
406-363-9202

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Hi Listers

I have for a long time been keen to find references to EMs in movies and
put them on a website for all to see. Every time I think about it I
promptly forget and the cycle repeats not this time!

I wondered is anyone has spotted EMs in movies/tv shows/documentaries?
If so what and where? If you can point out the make, model and any
inaccuracies all the better!

So far the ones I can remember are:

1. Flash Gordon 1936 (Controls of the ship are a TEM)
2. Independence Day in Area 51 scene
3. The Man in White Suit
4. Predator 2
5. CSI had a Hitachi S3400 once I recall
6. There was a fake one in Spiderman 2
7. Blade Runner

Any more would be greatly appreciated including those not in English!

Regards
John

.

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Quincy. In a big voice, "Let's put it in the scanning electron microscope."
I was told that the consultant was an ETEC customer. I always thought it
was great because I knew what Quincy was talking about (even if he didn't).

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: john.mitchels-at-gmail.com [mailto:john.mitchels-at-gmail.com]
Sent: Monday, January 23, 2012 5:01 PM
To: kenconverse-at-qualityimages.biz

Hi Listers

I have for a long time been keen to find references to EMs in movies and
put them on a website for all to see. Every time I think about it I
promptly forget and the cycle repeats not this time!

I wondered is anyone has spotted EMs in movies/tv shows/documentaries?
If so what and where? If you can point out the make, model and any
inaccuracies all the better!

So far the ones I can remember are:

1. Flash Gordon 1936 (Controls of the ship are a TEM)
2. Independence Day in Area 51 scene
3. The Man in White Suit
4. Predator 2
5. CSI had a Hitachi S3400 once I recall
6. There was a fake one in Spiderman 2
7. Blade Runner

Any more would be greatly appreciated including those not in English!

Regards
John

.

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So far thanks to over 60 responses (and a quick trawl of the archives),
this is great keep them coming!

We are up to the following:

1. Flash Gordon
2. Independence Day
3. The Man in White Suit
4. Predator 2
5. CSI
6. Spiderman 2
7. Blade Runner
8. NCIS
9. Dexter
10.Contagion
11. Andromeda Strain The Andromeda Strain (1971)and then the remake 2008
tv-miniseries.
12. Quincy
13. X-Files
14.Pres Obama at Intel's Titan
15.Spiderman 1
16.American Pickers
17.The Last Mimzy
18 ALTO MEDIA 'travel' into the matter.
19. special effects for one of the Star Trek
20. Batman
21. Jurassic Park
22. The Bone Collector" analyzes something
23. Avatar
24. The Prisoner (British TV series)
25. The Naked Gun

Thanks for all the responses so far! Once collected I will put together
a website and send the link to the list.
John

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Hello out there,

We have a Leo 1550 FEG-SEM that isn't running as well as it should be. We're not quite sure what the problem is exactly, but we know what the results are.

A few things are not fully functioning as our beam current is on average in the low 40 micro-amp range. Another issue is the alignment of the beam to the aperatures. When we select different apertures, the beam is supposed to be calibrated to the exact coordinates of the aperture and be deflected through the one selected. This is not happening though, instead we get a beam that moves slightly off from where it was originally, which in our case seems to be at the 30um aperture.

Does anyone out there have any experience to these columns and know of any kind of solution to these problems? I'm also looking for some sort of manual or user guide to the SEM, hopefully in pdf form, if anyone is willing to share.

Thank you for your time,

Andrew Ornelas�
Metallurgical Lab Technician
Intertek APTECH
601 West California Avenue�
Sunnyvale, CA 94086��

Tel. (Direct):�(408) 636-5308�
Tel. (Main):�(408) 745-7000�
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Valued Quality. Delivered.
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Is "The Man In The White Suit" the earliest scene with an EM in
movies that anyone knows of?

Great movie by the way.
john

At 02:01 PM 1/23/2012, you wrote:

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7, 24 -- From donovan-at-uoregon.edu Mon Jan 23 17:57:23 2012
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Hi,

I agree with everything Warren says, the average EDS/EDX system is not set up for boron analysis out of the box. You need to set it up yourself, and then test with known reference materials.

I believe the best way is with WDS EPMA (probe) analysis. Most probes have special diffracting crystals optimised for the boron x-ray.

cheers,
Ron
________________________________________
X-from: wesaia-at-iastate.edu [wesaia-at-iastate.edu]
Sent: Tuesday, 24 January 2012 3:20 AM
To: Ron Rasch

The first thing you do is proceed with great caution. If your system can detect boron, I am sure it will spit out a number. It might even have some resemblance to the truth, but probably not.

The second thing you should probably do is start looking into the difficulty of light element analysis. You have boron and oxygen (light elements) combined with zinc (a rather heavy element). Look into the ZAF factors and see how big the absorption corrections. Ask yourself how well you know the mass absorption coefficients for those combinations. See what you can do about changing conditions (dropping voltage) to reduce the magnitude of the corrections.

The third thing you do would be to make sure your system can see the boron peak. My guess is that you are probably not optimized to detect boron. We had an old Kevex Quantum detector running on one of the early IXRF systems. You might be able to adjust your discriminators to better see the peak. We had to. In the end, we could see most of the boron peak, but not much if any of the background on the low energy side.

The fourth thing would be to get some reference materials. If your system cannot give you right answers on a known compound (and it probably won't), then you can forget about trying to analyze an unknown compound. You may be able to collect your own standards to help the matter, but that is not a job for novices. (If you succeed, you may have progressed beyond the novice stage. There is much more to be said on this topic.)

Another thing is to consider the limitations of EDS in general. I suppose they will not have a flat, polished sample. You'll need to consider the effects of geometry. They affect results and become more significant the lighter the elements. Depending on where you hit a particle (on the side near the detector or the side away), you can probably produce any results you want.

Finally, I would get a large supply of salt so you can hand out your results with many grains. Seriously, I would try to get your client to accept qualitative results such as "The sample shows Zn, B, and O. It could be borate." This will be even more convincing if you can compare spectra of their unknown to a known sample of zinc borate and say, "See, they match". That may be as far as you really want to push it.

Regards,
Warren Straszheim

-----Original Message-----
X-from: donk-at-ardl.com [mailto:donk-at-ardl.com]
Sent: Monday, January 23, 2012 10:42 AM
To: wesaia-at-iastate.edu

"The Andromeda Strain (1971) featured a "Forgflo" TEM (formerly RCA EMU-3). Forgflo Corp. took over the RCA EM business around 1970, just in time to get the Forgflo name on a TEM in that movie. The movie was great, but of course the depiction of the use of the TEM was pretty ridiculous. Curiously, a few years ago I was chatting with a woman on a flight out of Tokyo Narita airport, while we were getting settled in the business cabin. When I mentioned working with electron microscopes, she said that her father, a long time ago, owned a company that for a short while sold EMs. She was amazed when I said "Oh, you mean Forgflo...the microscope in The Andromeda Strain"... :-). I've never personally seen a microscope with the Forgflo logo on it...I wonder if anyone on the listserver has any familiarity with the name...?

Also, "Avatar" has a giant machine in the early part of the movie that I believe is supposed to be an electron microscope...check it out.

Larry

Dr. Lawrence F. Allard, FMSA
Distinguished Research Staff Member
High Temperature Materials Laboratory
Microscopy Group
Materials Science and Technology Division
Oak Ridge National Laboratory
1 Bethel Valley Road
PO Box 2008
Oak Ridge, TN 37831-6064
(note: the last 4 lines are sufficient for mailing or overnight courier service)
865-607-1144 (cell)
865-576-5413 (fax)
allardLFjr-at-ornl.gov

-----Original Message-----
X-from: donovan-at-uoregon.edu [mailto:donovan-at-uoregon.edu]
Sent: Monday, January 23, 2012 6:58 PM
To: Allard Jr, Lawrence Frederick

Is "The Man In The White Suit" the earliest scene with an EM in
movies that anyone knows of?

Great movie by the way.
john

At 02:01 PM 1/23/2012, you wrote:

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Re to Bryan Hansen,

Hi All too,

this just to try to figure out whether you would like to get not only references but (if you like) some .pdf
on the making and the method used for staining.

Interesting method, but not the least for a substitution of uranyl-acetate (see below)

e.g. Abstract 1993:
27-V-1030 Enhancement of the BSE signal from hydrous SEM samples by use of a platinum blue.
Keiichi Tanaka and Kenji Inagaki
Seirei Christopher Coll. Nurs. ,Hamamatsu, 433 Japan.
The most serious problem for observing hydrous biological samples is deterioration of signal-noise ratio in consequence of lowering of vacuum in specimen chambers. Although metal coating is usually used for increasing SE and BSE yields on ordinary SEM studies, the method cannot be applied for wet samples. Instead of the metal coating method, therefore, we newly devised a staining method with a 'platinum blue'. Platinum blue is a general term of polymeric compounds of deep blue in which platinum [is?] coordinated to amide
groups. In this study, we used a platinum blue prepared from the reaction of cis-dichlorodiammineplatinum (II) with thymidine, being expected to have a molecular formula [Pt4 (NHx)x . (CxHxxOx) 4] +x.

{Note added by W. Muss: unfortunately the term [Pt4 (NHx)x . (CxHxxOx) 4] +x is printed so small (poor quality) that identification of Atom-numbers is not possible from this specific reprint, but from the paper out of 2007 - see below - it is ([Pt4(NH3)8(C6H13O5)4]^+5) pH = 3-4. }

This compound did not crystallize and the Pt particles were not recognized on specimen surfaces at the observation of 100.000X. The staining method was as follows. The specimens were dipped in the
concentrated solution for about 15-30 min and rinsed in distilled water. After removing excess fluid, they are immediately observed in a SEM for wet samples.
This method was very effective for enhancing SE and BSE yields, consequently, we could get enough signals for obtaining specimen images even at the vacuum of a 4.0 Torr.

or: Paper from 2007
"Paper introduces an aqueous solution of Platinum Blue (Pt-Blue) as an alternative to Uranyl actetate (UA) for staining in Transmission EM.....

There is also a paper from 2009..

On 7th of Dec. 2011 I posted the following in TOPICS/ResearchGate
(==} { http://www.researchgate.net/topic/Electron_Microscopy/ } ) or - better, since directly -

{ http://www.researchgate.net/topic/Electron_Microscopy/post/Edition_of_post_12-01-23_It_is_already_known_for_years_that_eg_lanthanum_nitrate_as_a_reagent_either_added_to_the_fixative_or_as_separate_incubation_solution_cave_specific_recipes_to_be_followed_has_s }

Header: (TEM): Substitute reagents for Uranyl-Acetate in positive and negative staining of resin ultrathin sections.

{Dear Colleagues,
I would like to inform you of an article which recently was published in Journal of Electron Microscopy (Toyko) which I found to be perhaps of interest also to you Electron Microscopists:

Masamichi Nakakoshi, Hideo Nishioka, and Eisaku Katayama
New versatile staining reagents for biological transmission electron microscopy that substitute for uranyl acetate}

Just only citing/referencing......not yet working with Platinum Blue,
but trying to play with the Lanthanides Samarium- & or Gadolinium-Triacetate-opportunity

Please tell me, whether you will accept sending of pdf's to your mail account or not...

Best regards,

Wolfgang MUSS
Member of MSA
SALZBURG, Austria

} -----Urspr�ngliche Nachricht-----
} Von: bryan.todd.hansen-at-gmail.com [mailto:bryan.todd.hansen-at-gmail.com]
} Gesendet: Montag, 23. J�nner 2012 22:29
} An: Mu� Wolfgang
} Betreff: [Microscopy] Platinum Blue Recipe
} ----------------------------------------------------------------------------
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}
} Hi All,
}
} I've found a couple of papers that list Pt-blue as an alternative to
} Uranyl
} Acetate, but I haven't been able to track down the way to make it.
} Does
} anyone have a working recipe as how to make the Pt-blue? Thanks for
} any
} help/tips.
}
} --
} Bryan Hansen
} EM Technician
} Rocky Mountain Laboratory
} NIAID/NIH
} bryan.todd.hansen-at-gmail.com
} hansenbry-at-niaid.nih.gov
} 406-363-9202
}
}
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I was watching the second episode of the first season of the english show
sherlock last night, Sherlock used a Leica dissecting microscope to get
some lovely scanning EM images of pollen which were characterized as a
pathogen�

Simon

Simon C. Watkins PhD
Professor, Cell Biology
Professor, Immunology
Vice Chair Cell Biology
Director Center for Biologic Imaging
University of Pittsburgh
BSTS 225

3500 Terrace St
Pittsburgh PA 15261
412-352-2277
Www.cbi.pitt.edu

On 1/23/12 10:51 PM, "allardlfjr-at-ornl.gov" {allardlfjr-at-ornl.gov} wrote:

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The Forgflo EM in The Andromeda Strain was a 4C model. Just as I was
finishing my Master's at Cal State Long Beach the Biology dept. bought
one. Our service engineer was the one who set up the EM for the movie.

Geoff

On 1/23/2012 10:48 PM, allardlfjr-at-ornl.gov wrote:
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} "The Andromeda Strain (1971) featured a "Forgflo" TEM (formerly RCA EMU-3). Forgflo Corp. took over the RCA EM business around 1970, just in time to get the Forgflo name on a TEM in that movie. The movie was great, but of course the depiction of the use of the TEM was pretty ridiculous. Curiously, a few years ago I was chatting with a woman on a flight out of Tokyo Narita airport, while we were getting settled in the business cabin. When I mentioned working with electron microscopes, she said that her father, a long time ago, owned a company that for a short while sold EMs. She was amazed when I said "Oh, you mean Forgflo...the microscope in The Andromeda Strain"... :-). I've never personally seen a microscope with the Forgflo logo on it...I wonder if anyone on the listserver has any familiarity with the name...?
}
} Also, "Avatar" has a giant machine in the early part of the movie that I believe is supposed to be an electron microscope...check it out.
}
} Larry
}
} Dr. Lawrence F. Allard, FMSA
} Distinguished Research Staff Member
} High Temperature Materials Laboratory
} Microscopy Group
} Materials Science and Technology Division
} Oak Ridge National Laboratory
} 1 Bethel Valley Road
} PO Box 2008
} Oak Ridge, TN 37831-6064
} (note: the last 4 lines are sufficient for mailing or overnight courier service)
} 865-607-1144 (cell)
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} allardLFjr-at-ornl.gov
}
} -----Original Message-----
} X-from: donovan-at-uoregon.edu [mailto:donovan-at-uoregon.edu]
} Sent: Monday, January 23, 2012 6:58 PM
} To: Allard Jr, Lawrence Frederick
} Subject: [Microscopy] Re: EMs in the Movies
}
}
}
}
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} Is "The Man In The White Suit" the earliest scene with an EM in
} movies that anyone knows of?
}
} Great movie by the way.
} john
}
} At 02:01 PM 1/23/2012, you wrote:
}
}
}
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} }
} } Hi Listers
} }
} } I have for a long time been keen to find references to EMs in movies and
} } put them on a website for all to see. Every time I think about it I
} } promptly forget and the cycle repeats not this time!
} }
} } I wondered is anyone has spotted EMs in movies/tv shows/documentaries?
} } If so what and where? If you can point out the make, model and any
} } inaccuracies all the better!
} }
} } So far the ones I can remember are:
} }
} } 1. Flash Gordon 1936 (Controls of the ship are a TEM)
} } 2. Independence Day in Area 51 scene
} } 3. The Man in White Suit
} } 4. Predator 2
} } 5. CSI had a Hitachi S3400 once I recall
} } 6. There was a fake one in Spiderman 2
} } 7. Blade Runner
} }
} } Any more would be greatly appreciated including those not in English!
} }
} } Regards
} } John
} }
} }
} } .
} }
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--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583
mcauliff-at-umdnj.edu
**********************************************

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Don't know how this topic came up, but of course Blade Runner has a
scene featuring a behind-the counter SEM with no need for sample prep,
used to quickly distinguish a scale of a replicant-snake from a
replicant-fish. Another scene features an image-enhancing device that
might in retrospect be seen as a form of light-field imaging.

Peter

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2, 19 -- From: Peter Werner {germpore-at-sonic.net}
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2, 19 -- Subject: Re: [Microscopy] EMs in the Movies
2, 19 -- Date: Tue, 24 Jan 2012 16:38:57 -0800
2, 19 -- References: {201201240355.q0O3t2WA022366-at-ns.microscopy.com}
2, 19 -- X-Mailer: Apple Mail (2.936)
==============================End of - Headers==============================

Contents Retrieved from Microscopy Listserver Archives
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You can watch the BBC on iPlayer if you can get a proxy server that makes it look like you are in
the UK. We just watched the second series of Sherlock.

Sent from my iPad.
--
John Mansfield PhD Cphys MInstP
2010 Microscopy & Microanalysis Program Chair
North Campus Electron Microbeam Analysis Laboratory
417 SRB, University of Michigan
2455 Hayward, Ann Arbor MI 48109-2143 USA
Phone: (734) 936-3352 FAX (734) 763-2282 Cell. Phone: (734) 834-3913

4304 Spring Lake Boulevard
Ann Arbor MI 48108-9657
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Please note: Electronic Mail is not secure, but should be read several times every day, and should
definitely be used for urgent or sensitive issues.

On Jan 24, 2012, at 9:00 AM, microscopylistserver-noreply-at-microscopy.com wrote:

}
}
}
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} I was watching the second episode of the first season of the english show
} sherlock last night, Sherlock used a Leica dissecting microscope to get
} some lovely scanning EM images of pollen which were characterized as a
} pathogen�
}
} Simon
}
} Simon C. Watkins PhD
} Professor, Cell Biology
} Professor, Immunology
} Vice Chair Cell Biology
} Director Center for Biologic Imaging
} University of Pittsburgh
} BSTS 225
}
} 3500 Terrace St
} Pittsburgh PA 15261
} 412-352-2277
} Www.cbi.pitt.edu
}
}
}
}
}
}
} On 1/23/12 10:51 PM, "allardlfjr-at-ornl.gov" {allardlfjr-at-ornl.gov} wrote:
}
} }
} }
} }
} } --------------------------------------------------------------------------
} } --
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
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} }
} } "The Andromeda Strain (1971) featured a "Forgflo" TEM (formerly RCA
} } EMU-3). Forgflo Corp. took over the RCA EM business around 1970, just in
} } time to get the Forgflo name on a TEM in that movie. The movie was
} } great, but of course the depiction of the use of the TEM was pretty
} } ridiculous. Curiously, a few years ago I was chatting with a woman on a
} } flight out of Tokyo Narita airport, while we were getting settled in the
} } business cabin. When I mentioned working with electron microscopes, she
} } said that her father, a long time ago, owned a company that for a short
} } while sold EMs. She was amazed when I said "Oh, you mean Forgflo...the
} } microscope in The Andromeda Strain"... :-). I've never personally seen a
} } microscope with the Forgflo logo on it...I wonder if anyone on the
} } listserver has any familiarity with the name...?
} }
} } Also, "Avatar" has a giant machine in the early part of the movie that I
} } believe is supposed to be an electron microscope...check it out.
} }
} } Larry
} }
} } Dr. Lawrence F. Allard, FMSA
} } Distinguished Research Staff Member
} } High Temperature Materials Laboratory
} } Microscopy Group
} } Materials Science and Technology Division
} } Oak Ridge National Laboratory
} } 1 Bethel Valley Road
} } PO Box 2008
} } Oak Ridge, TN 37831-6064
} } (note: the last 4 lines are sufficient for mailing or overnight courier
} } service)
} } 865-607-1144 (cell)
} } 865-576-5413 (fax)
} } allardLFjr-at-ornl.gov
} }
} } -----Original Message-----
} } X-from: donovan-at-uoregon.edu [mailto:donovan-at-uoregon.edu]
} } Sent: Monday, January 23, 2012 6:58 PM
} } To: Allard Jr, Lawrence Frederick
} } Subject: [Microscopy] Re: EMs in the Movies
} }
} }
} }
} }
} } --------------------------------------------------------------------------
} } --
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
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} } --------------------------------------------------------------------------
} } --
} }
} } Is "The Man In The White Suit" the earliest scene with an EM in
} } movies that anyone knows of?
} }
} } Great movie by the way.
} } john
} }
} } At 02:01 PM 1/23/2012, you wrote:
} }
} }
} }
} } } -------------------------------------------------------------------------
} } } ---
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } } America
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} } }
} } } Hi Listers
} } }
} } } I have for a long time been keen to find references to EMs in movies and
} } } put them on a website for all to see. Every time I think about it I
} } } promptly forget and the cycle repeats not this time!
} } }
} } } I wondered is anyone has spotted EMs in movies/tv shows/documentaries?
} } } If so what and where? If you can point out the make, model and any
} } } inaccuracies all the better!
} } }
} } } So far the ones I can remember are:
} } }
} } } 1. Flash Gordon 1936 (Controls of the ship are a TEM)
} } } 2. Independence Day in Area 51 scene
} } } 3. The Man in White Suit
} } } 4. Predator 2
} } } 5. CSI had a Hitachi S3400 once I recall
} } } 6. There was a fake one in Spiderman 2
} } } 7. Blade Runner
} } }
} } } Any more would be greatly appreciated including those not in English!
} } }
} } } Regards
} } } John
} } }
} } }
} } } .
} } }
} } } ==============================Original
} } } Headers==============================
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} } ==============================Original
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} } 17, 30 -- ([160.91.2.112]) with mapi; Mon, 23 Jan 2012 22:47:35 -0500
} } 17, 30 -- From: "Allard Jr, Lawrence Frederick" {allardlfjr-at-ornl.gov}
} } 17, 30 -- To: "'donovan-at-uoregon.edu'" {donovan-at-uoregon.edu} ,
} } 17, 30 -- "'microscopy-at-microscopy.com'"
} } {microscopy-at-microscopy.com}
} } 17, 30 -- Date: Mon, 23 Jan 2012 22:47:34 -0500
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}
}
}
}
} ==============================Original Headers==============================
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Contents Retrieved from Microscopy Listserver Archives
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A life science professor watched Sherlock movies. As a biological processor, he used
Light microscopes a lot. So he saw something to his profession. Like me, anything related To
electron microscopes, I would be very interested.

Zhen

Sent from my iPad

On Jan 24, 2012, at 7:13 AM, "microscopylistserver-noreply-at-microscopy.com"
{microscopylistserver-noreply-at-microscopy.com} wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} I was watching the second episode of the first season of the english show
} sherlock last night, Sherlock used a Leica dissecting microscope to get
} some lovely scanning EM images of pollen which were characterized as a
} pathogenŠ
}
} Simon
}
} Simon C. Watkins PhD
} Professor, Cell Biology
} Professor, Immunology
} Vice Chair Cell Biology
} Director Center for Biologic Imaging
} University of Pittsburgh
} BSTS 225
}
} 3500 Terrace St
} Pittsburgh PA 15261
} 412-352-2277
} Www.cbi.pitt.edu
}
}
}
}
}
}
} On 1/23/12 10:51 PM, "allardlfjr-at-ornl.gov" {allardlfjr-at-ornl.gov} wrote:
}
} }
} }
} }
} } --------------------------------------------------------------------------
} } --
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } --------------------------------------------------------------------------
} } --
} }
} } "The Andromeda Strain (1971) featured a "Forgflo" TEM (formerly RCA
} } EMU-3). Forgflo Corp. took over the RCA EM business around 1970, just in
} } time to get the Forgflo name on a TEM in that movie. The movie was
} } great, but of course the depiction of the use of the TEM was pretty
} } ridiculous. Curiously, a few years ago I was chatting with a woman on a
} } flight out of Tokyo Narita airport, while we were getting settled in the
} } business cabin. When I mentioned working with electron microscopes, she
} } said that her father, a long time ago, owned a company that for a short
} } while sold EMs. She was amazed when I said "Oh, you mean Forgflo...the
} } microscope in The Andromeda Strain"... :-). I've never personally seen a
} } microscope with the Forgflo logo on it...I wonder if anyone on the
} } listserver has any familiarity with the name...?
} }
} } Also, "Avatar" has a giant machine in the early part of the movie that I
} } believe is supposed to be an electron microscope...check it out.
} }
} } Larry
} }
} } Dr. Lawrence F. Allard, FMSA
} } Distinguished Research Staff Member
} } High Temperature Materials Laboratory
} } Microscopy Group
} } Materials Science and Technology Division
} } Oak Ridge National Laboratory
} } 1 Bethel Valley Road
} } PO Box 2008
} } Oak Ridge, TN 37831-6064
} } (note: the last 4 lines are sufficient for mailing or overnight courier
} } service)
} } 865-607-1144 (cell)
} } 865-576-5413 (fax)
} } allardLFjr-at-ornl.gov
} }
} } -----Original Message-----
} } X-from: donovan-at-uoregon.edu [mailto:donovan-at-uoregon.edu]
} } Sent: Monday, January 23, 2012 6:58 PM
} } To: Allard Jr, Lawrence Frederick
} } Subject: [Microscopy] Re: EMs in the Movies
} }
} }
} }
} }
} } --------------------------------------------------------------------------
} } --
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } --------------------------------------------------------------------------
} } --
} }
} } Is "The Man In The White Suit" the earliest scene with an EM in
} } movies that anyone knows of?
} }
} } Great movie by the way.
} } john
} }
} } At 02:01 PM 1/23/2012, you wrote:
} }
} }
} }
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} } }
} } } Hi Listers
} } }
} } } I have for a long time been keen to find references to EMs in movies and
} } } put them on a website for all to see. Every time I think about it I
} } } promptly forget and the cycle repeats not this time!
} } }
} } } I wondered is anyone has spotted EMs in movies/tv shows/documentaries?
} } } If so what and where? If you can point out the make, model and any
} } } inaccuracies all the better!
} } }
} } } So far the ones I can remember are:
} } }
} } } 1. Flash Gordon 1936 (Controls of the ship are a TEM)
} } } 2. Independence Day in Area 51 scene
} } } 3. The Man in White Suit
} } } 4. Predator 2
} } } 5. CSI had a Hitachi S3400 once I recall
} } } 6. There was a fake one in Spiderman 2
} } } 7. Blade Runner
} } }
} } } Any more would be greatly appreciated including those not in English!
} } }
} } } Regards
} } } John
} } }
} } }
} } } .
} } }
} } } ==============================Original
} } } Headers==============================
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Our "Almost Famous" TEM:

Several of you mentioned Blade Runner, directed by Ridley Scott.

Back in 1994, we were contacted by an assistant to Mr. Scott for lease
of our recently purchased, Hitachi H7100 TEM. They were completing
construction of a laboratory set for the film, "Crisis in The Hot
Zone," and Mr. Scott wanted a functioning TEM, as was described in the
book by Richard Preston. The film was to star Robert Redford and Jodie
Foster (who was to operate the TEM).

The production assistant contacted Hitachi to pursue the lease of a
TEM for the interior shots, but Hitachi informed them the TEM they
wanted was to be delivered to SIU that month. After speaking to the
assistant, we agreed to have the TEM delivered to the set in Los
Angeles, where HItachi technicians would install and make the scope
operational for filming. Afterwards, it would be repacked and shipped
to SIU for regular installation. We had already dubbed it, The Jodie
Foster Microscope………. sigh.

Several weeks went by and we received a FAX from the assistant
indicating that the film was being shelved due to script disagreements
and since a similar film, Outbreak, was to be released at about the
same time. I saw the awful film and was quite disappointed with the
science portrayed in the film. Outbreak was fictional; whereas, Hot
Zone is based on an actual incident. Steven King is said to have
commented that Hot Zone as the scariest thing he ever read.

Supposedly, a more recent film, "Contagion." with Gwyneth Paltrow and
Matt Damon "feels very much like the movie that should have been
adapted from the Hot Zone. Has anyone seen "Contagion"? I don't think
it did well in the box office. Any TEM's in that film?

Cheers,

--
John J. Bozzola, Ph.D., Professor & Director of IMAGE (Happily Retired :-)
Integrated Microscopy & Graphics Expertise
Southern Illinois University
750 Communications Drive
Carbondale, IL 62901
Phone: 618-453-3730

===========================================================

} Don't know how this topic came up, but of course Blade Runner has a
} scene featuring a behind-the counter SEM with no need for sample prep,
} used to quickly distinguish a scale of a replicant-snake from a
} replicant-fish. Another scene features an image-enhancing device that
} might in retrospect be seen as a form of light-field imaging.
}
} Peter
}
} ==============================Original

==============================Original Headers==============================
11, 21 -- From bozzola-at-siu.edu Tue Jan 24 21:33:51 2012
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11, 21 -- Date: Tue, 24 Jan 2012 21:33:50 -0600
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11, 21 -- Subject: Re: [Microscopy] Re: EMs in the Movies
11, 21 -- From: John Bozzola {bozzola-at-siu.edu}
11, 21 -- To: MSAListserver {Microscopy-at-microscopy.com}
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Contents Retrieved from Microscopy Listserver Archives
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Dear "happily retired" John,

Contagion is on the list that was sent out. It features and FEI Titan. However, I hear that the company only shipped the outer box for filming. It looks like Gwyneth Paltrow never was able to use it!

If we are looking at microscopes on screen, our FEI G2 20 was featured in a TV advert for the University of Phoenix. We had the film crew here for a day. I estimated about 45 people, all the equipment you ever would need for any sort of scene, and a whole day of filming. The shot on the screen was about 2 seconds long.

Working in Los Angels, we sometimes get good deals on used equipment. For example, we have a pink Zeiss light microscope that featured in the first Jurassic Park movie. Apparently pink objects look white on film, so the machine was painted pink.

The "biggest electron microscope on the East Coast" featured in the first "Spiderman" (and where Peter Parker was bitten by the spider) was a model constructed around a statue in the front atrium of the LA Natural History Museum. Is there anyone on the list from the museum? It would be great to get another "back-stage" look at the collection.

The SEM used by Harrison Ford in Blade Runner was supposedly located in the Los Angeles Chinatown. We guessed it took place in the future because it was raining.

Best wishes on your retirement John.

Paul.

X-from Still Sunny California
House Research Institute (we changed our name!)
2100 W 3rd St
Los Angeles
CA 90057.

-----Original Message-----
X-from: bozzola-at-siu.edu [mailto:bozzola-at-siu.edu]
Sent: Tue 1/24/2012 7:36 PM
To: Webster, Paul

Our "Almost Famous" TEM:

Several of you mentioned Blade Runner, directed by Ridley Scott.

Back in 1994, we were contacted by an assistant to Mr. Scott for lease
of our recently purchased, Hitachi H7100 TEM. They were completing
construction of a laboratory set for the film, "Crisis in The Hot
Zone," and Mr. Scott wanted a functioning TEM, as was described in the
book by Richard Preston. The film was to star Robert Redford and Jodie
Foster (who was to operate the TEM).

The production assistant contacted Hitachi to pursue the lease of a
TEM for the interior shots, but Hitachi informed them the TEM they
wanted was to be delivered to SIU that month. After speaking to the
assistant, we agreed to have the TEM delivered to the set in Los
Angeles, where HItachi technicians would install and make the scope
operational for filming. Afterwards, it would be repacked and shipped
to SIU for regular installation. We had already dubbed it, The Jodie
Foster Microscope�?��?��?�. sigh.

Several weeks went by and we received a FAX from the assistant
indicating that the film was being shelved due to script disagreements
and since a similar film, Outbreak, was to be released at about the
same time. I saw the awful film and was quite disappointed with the
science portrayed in the film. Outbreak was fictional; whereas, Hot
Zone is based on an actual incident. Steven King is said to have
commented that Hot Zone as the scariest thing he ever read.

Supposedly, a more recent film, "Contagion." with Gwyneth Paltrow and
Matt Damon "feels very much like the movie that should have been
adapted from the Hot Zone. Has anyone seen "Contagion"? I don't think
it did well in the box office. Any TEM's in that film?

Cheers,

--
John J. Bozzola, Ph.D., Professor & Director of IMAGE (Happily Retired :-)
Integrated Microscopy & Graphics Expertise
Southern Illinois University
750 Communications Drive
Carbondale, IL 62901
Phone: 618-453-3730

===========================================================

} Don't know how this topic came up, but of course Blade Runner has a
} scene featuring a behind-the counter SEM with no need for sample prep,
} used to quickly distinguish a scale of a replicant-snake from a
} replicant-fish. Another scene features an image-enhancing device that
} might in retrospect be seen as a form of light-field imaging.
}
} Peter
}
} ==============================Original

==============================Original Headers==============================
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My favourite example is from the X-Files. Scully visits a lab and watches the electron microscopist use a TEM. She then gives Scully an SEM image of a pollen grain or spore and tells her it must be the alien virus. Needless to say, the electron microscopist is involved in a fatal automobile accident that evening.

Dave

-----Original Message-----
X-from: David Patton
Sent: 24 January 2012 16:50
To: 'john.mitchels-at-gmail.com'

So far thanks to over 60 responses (and a quick trawl of the archives),
this is great keep them coming!

We are up to the following:

1. Flash Gordon
2. Independence Day
3. The Man in White Suit
4. Predator 2
5. CSI
6. Spiderman 2
7. Blade Runner
8. NCIS
9. Dexter
10.Contagion
11. Andromeda Strain The Andromeda Strain (1971)and then the remake 2008
tv-miniseries.
12. Quincy
13. X-Files
14.Pres Obama at Intel's Titan
15.Spiderman 1
16.American Pickers
17.The Last Mimzy
18 ALTO MEDIA 'travel' into the matter.
19. special effects for one of the Star Trek
20. Batman
21. Jurassic Park
22. The Bone Collector" analyzes something
23. Avatar
24. The Prisoner (British TV series)
25. The Naked Gun

Thanks for all the responses so far! Once collected I will put together
a website and send the link to the list.
John

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Dear List,
dear John,

out of my files, just as a supplement to your posts:

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There is also an orange column TEM in the TV series {Dexter} . When he's in his little lab.
I too have been bored by TV and movies.

More TEM's in movies I say.... actually how about an old VG HB501. I know the one here has more character and versatility than some leading actors and actresses out there and it works for free.
The microscopist on the other hand will need food, drink and board.
{ { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { { {

best regards and wishes,

Wolfgang MUSS
SALZBURG-AUSTRIA

----

} -----Urspr�ngliche Nachricht-----
} Von: john.mitchels-at-gmail.com [mailto:john.mitchels-at-gmail.com]
} Gesendet: Montag, 23. J�nner 2012 23:01
} An: Mu� Wolfgang
} Betreff: [Microscopy] EMs in the Movies
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
} Hi Listers
}
} I have for a long time been keen to find references to EMs in movies
} and put them on a website for all to see. Every time I think about it I
} promptly forget and the cycle repeats not this time!
}
} I wondered if anyone has spotted EMs in movies/tv shows/documentaries?
}
} If so what and where? If you can point out the make, model and any
} inaccuracies all the better!
}
} So far the ones I can remember are:
}
} 1. Flash Gordon 1936 (Controls of the ship are a TEM)
} 2. Independence Day in Area 51 scene
} 3. The Man in White Suit
} 4. Predator 2
} 5. CSI had a Hitachi S3400 once I recall
} 6. There was a fake one in Spiderman 2
} 7. Blade Runner
}
} Any more would be greatly appreciated including those not in English!
}
} Regards
} John
}
}
} .
}
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What I take from this story is that University of Phoenix wants to
portray that they have electron microscopes, yet they have no such
thing at any of their many campuses.

Jay Campbell

---------- Forwarded message ----------
X-from: PWebster-at-hei.org

Dear "happily retired" John,

Contagion is on the list that was sent out. It features and FEI Titan.
However, I hear that the company only shipped the outer box for
filming. It looks like Gwyneth Paltrow never was able to use it!

If we are looking at microscopes on screen, our FEI G2 20 was featured
in a TV advert for the University of Phoenix. We had the film crew
here for a day. I estimated about 45 people, all the equipment you
ever would need for any sort of scene, and a whole day of filming. The
shot on the screen was about 2 seconds long.

Working in Los Angels, we sometimes get good deals on used equipment.
For example, we have a pink Zeiss light microscope that featured in
the first Jurassic Park movie. Apparently pink objects look white on
film, so the machine was painted pink.

The "biggest electron microscope on the East Coast" featured in the
first "Spiderman" (and where Peter Parker was bitten by the spider)
was a model constructed around a statue in the front atrium of the LA
Natural History Museum. Is there anyone on the list from the museum?
It would be great to get another "back-stage" look at the collection.

The SEM used by Harrison Ford in Blade Runner was supposedly located
in the Los Angeles Chinatown. We guessed it took place in the future
because it was raining.

Best wishes on your retirement John.

Paul.

X-from Still Sunny California
House Research Institute (we changed our name!)
2100 W 3rd St
Los Angeles
CA 90057.

-----Original Message-----
X-from: bozzola-at-siu.edu [mailto:bozzola-at-siu.edu]
Sent: Tue 1/24/2012 7:36 PM
To: Webster, Paul

Our "Almost Famous" TEM:

Several of you mentioned Blade Runner, directed by Ridley Scott.

Back in 1994, we were contacted by an assistant to Mr. Scott for lease
of our recently purchased, Hitachi H7100 TEM. They were completing
construction of a laboratory set for the film, "Crisis in The Hot
Zone," and Mr. Scott wanted a functioning TEM, as was described in the
book by Richard Preston. The film was to star Robert Redford and Jodie
Foster (who was to operate the TEM).

The production assistant contacted Hitachi to pursue the lease of a
TEM for the interior shots, but Hitachi informed them the TEM they
wanted was to be delivered to SIU that month. After speaking to the
assistant, we agreed to have the TEM delivered to the set in Los
Angeles, where HItachi technicians would install and make the scope
operational for filming. Afterwards, it would be repacked and shipped
to SIU for regular installation. We had already dubbed it, The Jodie
Foster Microscopeā?¦ā?¦ā?¦. sigh.

Several weeks went by and we received a FAX from the assistant
indicating that the film was being shelved due to script disagreements
and since a similar film, Outbreak, was to be released at about the
same time. I saw the awful film and was quite disappointed with the
science portrayed in the film. Outbreak was fictional; whereas, Hot
Zone is based on an actual incident. Steven King is said to have
commented that Hot Zone as the scariest thing he ever read.

Supposedly, a more recent film, "Contagion." with Gwyneth Paltrow and
Matt Damon "feels very much like the movie that should have been
adapted from the Hot Zone. Has anyone seen "Contagion"? I don't think
it did well in the box office. Any TEM's in that film?

Cheers,

--
John J. Bozzola, Ph.D., Professor & Director of IMAGE (Happily Retired :-)
Integrated Microscopy & Graphics Expertise
Southern Illinois University
750 Communications Drive
Carbondale, ILĀ 62901
Phone: 618-453-3730

===========================================================

} Don't know how this topic came up, but of course Blade Runner has a
} scene featuring a behind-the counter SEM with no need for sample prep,
} used to quickly distinguish a scale of a replicant-snake from a
} replicant-fish. Another scene features an image-enhancing device that
} might in retrospect be seen as a form of light-field imaging.
}
} Peter
}
} ==============================Original

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What episode of American Pickers had an EM in it?

} So far thanks to over 60 responses (and a quick trawl of the archives),
} this is great keep them coming!
}
} We are up to the following:
}
} 1. Flash Gordon
} 2. Independence Day
} 3. The Man in White Suit
} 4. Predator 2
} 5. CSI
} 6. Spiderman 2
} 7. Blade Runner
} 8. NCIS
} 9. Dexter
} 10.Contagion
} 11. Andromeda Strain The Andromeda Strain (1971)and then the remake 2008
} tv-miniseries.
} 12. Quincy
} 13. X-Files
} 14.Pres Obama at Intel's Titan
} 15.Spiderman 1
} 16.American Pickers
} 17.The Last Mimzy
} 18 ALTO MEDIA 'travel' into the matter.
} 19. special effects for one of the Star Trek
} 20. Batman
} 21. Jurassic Park
} 22. The Bone Collector" analyzes something
} 23. Avatar
} 24. The Prisoner (British TV series)
} 25. The Naked Gun
}
}
} Thanks for all the responses so far! Once collected I will put together
} a website and send the link to the list.
} John

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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3, 26 -- Subject: Re: [Microscopy] Updated Movie List
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Doctor Who and the Hand of Fear - it's used to examine a stone hand (that
comes to life). Not sure of the make, but could be Cambridge
Instruments... I'll dig out the DVD.

Great idea,

Austin

P.S. Partial side view here

http://2.bp.blogspot.com/_ZNUdpril8U4/TJbIfCxZatI/AAAAAAAAC2k/kvYB7CMhYRU/s1
600/vlcsnap-2010-09-18-13h31m33s59.jpg

http://2.bp.blogspot.com/_ZNUdpril8U4/TJbIYub7DzI/AAAAAAAAC2c/S7HVZJAXusw/s1
600/vlcsnap-2010-09-18-13h11m01s37.jpg

http://classicalgallifrey.blogspot.com/2010/09/serial-87-hand-of-fear.html

} } -----Original Message-----
} } From: oshel1pe-at-cmich.edu [mailto:oshel1pe-at-cmich.edu]
} } Sent: 25 January 2012 16:47
} } To: AuntDaisy-at-gmail.com
} } Subject: [SPAM] [Microscopy] Re: Updated Movie List
} }
} }
} }
} }
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} --
} } --
} }
} } What episode of American Pickers had an EM in it?
} }
} } } So far thanks to over 60 responses (and a quick trawl of the archives),
} } } this is great keep them coming!
} } }
} } } We are up to the following:
} } }
} } } 1. Flash Gordon
} } } 2. Independence Day
} } } 3. The Man in White Suit
} } } 4. Predator 2
} } } 5. CSI
} } } 6. Spiderman 2
} } } 7. Blade Runner
} } } 8. NCIS
} } } 9. Dexter
} } } 10.Contagion
} } } 11. Andromeda Strain The Andromeda Strain (1971)and then the remake
} 2008
} } } tv-miniseries.
} } } 12. Quincy
} } } 13. X-Files
} } } 14.Pres Obama at Intel's Titan
} } } 15.Spiderman 1
} } } 16.American Pickers
} } } 17.The Last Mimzy
} } } 18 ALTO MEDIA 'travel' into the matter.
} } } 19. special effects for one of the Star Trek
} } } 20. Batman
} } } 21. Jurassic Park
} } } 22. The Bone Collector" analyzes something
} } } 23. Avatar
} } } 24. The Prisoner (British TV series)
} } } 25. The Naked Gun
} } }
} } }
} } } Thanks for all the responses so far! Once collected I will put together
} } } a website and send the link to the list.
} } } John
} }
} } --
} } Philip Oshel
} } Microscopy Facility Supervisor
} } Biology Department
} } 024C Brooks Hall
} } Central Michigan University
} } Mt. Pleasant, MI 48859
} } (989) 774-3576
} }
} } ==============================Original
} } Headers==============================
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} } 3, 26 -- To: {Microscopy-at-microscopy.com}
} } 3, 26 -- From: Philip Oshel {oshel1pe-at-cmich.edu}
} } 3, 26 -- Subject: Re: [Microscopy] Updated Movie List
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Hi Johnson!
�
The use of cacodylate as a buffer for EM is more a question of conservationism than anything else.
We don't use it because it is the best, we use it because it has been used successfully for more than 50 years and we are too lazy to try something else.
Some buffers are better for specific applications, but cacodylate works most of the time for a large panel of samples.
Contrary to phosphate buffers, cacodylate does not precipitate in presence of calcium. This is important because it has been shown that calcium stabilizes somes structures during fixation. Tris is not an option because it has an amine and that is what aldehydes react with.
Hepes is really a possible alternative, it is a strong buffer and it is compatible with calcium.
Personally I go with Hepes and I still have to see a bad fixation due to the buffer (which doesn't mean it is best for all samples of course).
�
Cheers,
Stephane
�
�
�
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Sent: Thursday, January 26, 2012 4:23 AM

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Email: reganhll-at-gmail.com Name: Johnson

Organization: FMLS

Title-Subject: [Filtered] EM

Message: Dear Listserv members,
It would be nice to know what are the advantages of Sodium Cacodylet buffer over other buffer. I
looked for literature but most of the books and documents speak about the safety aspects.
Greetings,
Johnson.

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Hello!
I am responsible for a FEI Tecnai G20 which was installed in 2005.
As we discussed several times in the list, it is advised to store computer parts for future replacement needs because the computers evolve a lot during the lifetime of a TEM (or was it meant to buy a complete second PC?).
I suppose that the interface cards come from FEI themselves, so they will probably be able to provide them for some time.
I am not really a genius in computer matters and I would need you opinion about which hardware part of the computer we need to store?
Many thanks in advance.

Stephane

==============================Original Headers==============================
2, 34 -- From nizets2-at-yahoo.com Thu Jan 26 04:48:36 2012
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2, 34 -- From: Stephane Nizet {nizets2-at-yahoo.com}
2, 34 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com}
2, 34 -- Subject: which computer parts to store?
2, 34 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
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Computers are fairly inexpensive (compared to the equipment they run).
I would have at least one complete computer standing by in the event
of failure.

In my experience, hard drives are the first to go, so back up your
hard drives from time to time and have a spare main HD ready to
replace the one in the computer.

John Bozzola

On Thu, Jan 26, 2012 at 4:49 AM, {nizets2-at-yahoo.com} wrote:
}
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}
} ----------------------------------------------------------------------------
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}
} Hello!
} I am responsible for a FEI Tecnai G20 which was installed in 2005.
} As we discussed several times in the list, it is advised to store computer parts for future replacement needs because the computers evolve a lot during the lifetime of a TEM (or was it meant to buy a complete second PC?).
} I suppose that the interface cards come from FEI themselves, so they will probably be able to provide them for some time.
} I am not really a genius in computer matters and I would need you opinion about which hardware part of the computer we need to store?
} Many thanks in advance.
}
} Stephane
}
} ==============================Original Headers==============================
} 2, 34 -- From nizets2-at-yahoo.com Thu Jan 26 04:48:36 2012
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} 2, 34 -- Date: Thu, 26 Jan 2012 02:48:35 -0800 (PST)
} 2, 34 -- From: Stephane Nizet {nizets2-at-yahoo.com}
} 2, 34 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com}
} 2, 34 -- Subject: which computer parts to store?
} 2, 34 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
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--
John J. Bozzola, Ph.D., Retired
Professor & Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

==============================Original Headers==============================
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8, 20 -- Subject: Re: [Microscopy] which computer parts to store?
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We have just upgraded hardware computer links and the Windows operating system on a Hitachi S3000N SEM.

The problems will not be same for an FEI except that any interface units used in either the Hitachi or FEI will have been built to a particular PC standard and these change. In terms of maintaining PC hardware therefore it would be worth looking into exactly what FEI boards will link with on a PC interface as well as any limitations dictated by operating systems. It may for instance only be feasible to link FEI boards with an XP system. But the real killer will be the interface which for internal boards was ISA, became EISA and is generally PCI now where even external systems have changed from RS232, various parallel ports, USB (1,2 & 3) and Firewire.

What I'm saying is that you need to know what is the most recent type of PC, operating system and interface that is compatible with the FEI boards and make sure you have a fully functioning system. It's surprising how quickly standards change in PCs and no amount of emulators and patched systems may be able to rescue you when all the old systems have gone.

Incidentally our S3000N now runs on Windows XP which is a vaste improvement on the old Windows NT4.0 system but it's unlikely that we will ever be able to update/upgrade further. Anyway good luck.

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: bozzola-at-siu.edu [bozzola-at-siu.edu]
Sent: 26 January 2012 11:47
To: Malcolm Haswell

Computers are fairly inexpensive (compared to the equipment they run).
I would have at least one complete computer standing by in the event
of failure.

In my experience, hard drives are the first to go, so back up your
hard drives from time to time and have a spare main HD ready to
replace the one in the computer.

John Bozzola

On Thu, Jan 26, 2012 at 4:49 AM, {nizets2-at-yahoo.com} wrote:
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hello!
} I am responsible for a FEI Tecnai G20 which was installed in 2005.
} As we discussed several times in the list, it is advised to store computer parts for future replacement needs because the computers evolve a lot during the lifetime of a TEM (or was it meant to buy a complete second PC?).
} I suppose that the interface cards come from FEI themselves, so they will probably be able to provide them for some time.
} I am not really a genius in computer matters and I would need you opinion about which hardware part of the computer we need to store?
} Many thanks in advance.
}
} Stephane
}
} ==============================Original Headers==============================
} 2, 34 -- From nizets2-at-yahoo.com Thu Jan 26 04:48:36 2012
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} 2, 34 -- Date: Thu, 26 Jan 2012 02:48:35 -0800 (PST)
} 2, 34 -- From: Stephane Nizet {nizets2-at-yahoo.com}
} 2, 34 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com}
} 2, 34 -- Subject: which computer parts to store?
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--
John J. Bozzola, Ph.D., Retired
Professor & Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

==============================Original Headers==============================
8, 20 -- From bozzola-at-siu.edu Thu Jan 26 05:39:04 2012
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In my experience with various instruments (microscopes and otherwise),
host computer parts most likely to fail (in rough order of likelihood) are:

1. Hard drives
2. Power supplies
3. Motherboards

As the computer ages, all these parts can be had for next to nothing, if
you keep your eye out for them on eBay, second hand shops, and even the
dumpster. Having a pile of working hard drives that can replace the
working one is a great comfort. Likewise power supplies and
motherboards, although I'm lucky enough to have a local repair shop that
has been able to repair both in several cases for a very reasonable
price. Most often mobos fail either because the CMOS battery dies, or
some of the capacitors fail, which is often indicated by a swollen
appearance, particularly at the top. These are relatively easy to
replace for someone who is handy with a soldering iron. Power supplies
are also easily and inexpensively repaired by those who know how.

Having just done this yesterday, the longevity of a computer can be
extended by a yearly cleaning of the inside of the computer, esp. fans
and heatsinks. I've seen some neglected computers that were *completely*
full of dust bunnies and running hotter than a two dollar pistol. Not
good for any of the components! I believe regular cleaning is one reason
that I'm still running an HP Vectra computer on our EDS system that just
had its 12th birthday (knock wood).

Jim

--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

Intaxication: Euphoria at getting a
tax refund, which lasts until you realize
that it was your money to start with.

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Generally, sodium cacodylate seems to extract less cytoplasm than
phosphate buffers. Advantage is more cytoplasm, disadvantage is less
contrast because there is more cytoplasm. There may well be (likely
are) specific procedures or specimens where cacodylate buffer is
better.
The arsenic content is likely more important.
You might want to have a look at "Biomedical Electron Microscopy" by
Maunsbach and Afzelius - they show many EMs comparing different
preparatory methods (fixatives, buffers, etc.), embedding resins, and
so on. A good book to have around.

Phil

} Message: Dear Listserv members,
} It would be nice to know what are the advantages of Sodium Cacodylet
} buffer over other buffer. I
} looked for literature but most of the books and documents speak
} about the safety aspects.
} Greetings,
} Johnson.

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Hi Stephane,

As others already suggested, first priority is to have:

1) Harddrive with "carbon copy" of currently running software - it is
very easy to create with DD (Disk Dump) utility;
2) Motherboard;
3) Processor;

But do not assume that interface cards in the PC are made by OEM of the
TEM. I am not familiar with Technai specifically, but chances are that
most (if not all) cards installed in the control computer are
off-the-shelf products. If such adapted card becomes unavailable from
its OEM, it will becom impossible (or next to impossible) to get from
the FEI. So by all means solicit help from someone who is very familiar
with PC hardware and get all the cards sitting on PC bus.

Cheers :)
Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 1/26/2012 5:49 AM, nizets2-at-yahoo.com wrote:
} ----------------------------------------------------------------------------
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}
} Hello!
} I am responsible for a FEI Tecnai G20 which was installed in 2005.
} As we discussed several times in the list, it is advised to store computer parts for future replacement needs because the computers evolve a lot during the lifetime of a TEM (or was it meant to buy a complete second PC?).
} I suppose that the interface cards come from FEI themselves, so they will probably be able to provide them for some time.
} I am not really a genius in computer matters and I would need you opinion about which hardware part of the computer we need to store?
} Many thanks in advance.
}
} Stephane
}
} ==============================Original Headers==============================
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Dear Stephane
As John Bozzola wisely recommends, have at least one complete computer
standing by with a ghost HD. Don't be sure that company will keep any spare
cards for a long time. I have no experience with FEI but JEOL was unable to
provide a spare graphic card just 7 years after the model (JSM5600LV)
appeared. Make sure your spare computer is in a working condition by putting
it on from time to time. This frenzy of consumarism wlll have no end and we
have to get prepared for the worse..
Best -yorgos

Dr Yorgos Nikas
Athens Innovative Microscopy
Skra 36 Voula 16673 GREECE

Tel/fax +30 210 8957677
mobile +30 6945 107477
eikonika-at-otenet.gr
www.aim.cat
***********************************

----- Original Message -----
X-from: {nizets2-at-yahoo.com}
To: {eikonika-at-otenet.gr}
Sent: Thursday, January 26, 2012 12:54 PM

Stephane raised a great question and there have been several great
suggestions. We have a lab full of older FEI SEM and TEM microscopes
and have discussed this with our field service engineer. Our engineer
tells us that FEI learned from the problems they had with the
availability of components in early computer-controlled microscopes
because so much functionality was on the microscope PC. I think FEI
was not unique in encountering this problem. The engineer told us that
FEI made a concerted effort to move as much functionality as possible
into the microscope and to use multiple fast ethernet connections to
communicate with the PC. This makes upgrading the PC easier because
fewer custom boards are used. We have an older dual-beam FIB where
component availability has been a real issue. We always get scrutiny
from our IS folks because we cannot move out of unsupported operating
systems...

I would have a frank conversation with FEI's service specialist about
where your particular microscope is on this development timeline and
what your particular exposure is to part availability. At a minimum, I
would always have a spare hard drive with a disk image of a working
system configured the way you want sitting in a sealed container in
the lab. This has saved our bacon many times...

Best regards,
John Minter
Eastman Kodak

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Dear All,
I am just in my first experience of doing wet cryo ultramicrotome on
diamond sectioning of a PE compound containg ca. 1um size particles.
First I tried dry sectioning on a glass knife at -120�C but the cuts
came down rolled up very small in diameter. I had no idea how to smooth
them. Cutting seems impossible below 100 nm.
Wet cryo cuts now are coming off in 70% DMSO at -50�C at least somewhat
flat but very compressed. Thickness cutting is now 50 nm.
Can you give my some basic advice what I can improve (might be a lot ;-) ).

Best wishes,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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Course Reminder

Quantifying immunogold label on EM thin sections

April 5th to 7th, 2012

House Research Institute, Los Angeles, CA 90057

This is a 3-day course aimed at providing instruction on how to qualify
immunogold label from images collected using transmission electron
microscopy.

During the past decade, new ways of quantifying gold labeling within cells
have been devised. Their efficiency and validity rely on sound principles of
specimen sampling, event counting and inferential statistics. These
stereological tools will be introduced in lectures as well as by using
practical examples. Participants will be provided with practical exercises
to illustrate gold counting methods, which can be taken away for future
reference.

This is an intensive, hands-on course that is organized by an expert
electron microscopist and immunologist, and taught by the developer of many
of these stereological tools, who is also an expert stereologist.

For further details contact:
Paul Webster, Ph.D.
House Research Institute
2100 West Third Street
Los Angeles
CA 90057

Or check out http://immunogold.houseresearch.org

==============================Original Headers==============================
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The Department of Materials at the University of Oxford (UK) seeks a specialist facility technical manager of the highest calibre with the ability to provide strategic leadership for a large, world-class electron microscope facility.

Please follow the link: https://www.recruit.ox.ac.uk/pls/hrisliverecruit/erq_jobspec_version_4.jobspec?p_id=101946 �for more information

Closing date for applications is 24th February.

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I don't know if this was mentioned earlier: My favorite EM in recent movies
is in "Invasion" with Nicole Kidman (2007). She plays a doctor trying to
figure out a virus, and they use an FEI TEM to look at the virus. The
interesting part is that when they move the digital camera in (side-mount),
the next scene shows ... color images of red blood cells swimming in a
liquid (blood) and being attacked by the virus. I guess you can call that
"poetic license". I am certainly not claiming that our cameras (which they
use) can do that :-)

Mike Bode, Ph.D.
ResAlta Research Technologies Corp.

2102 Beech Ct
Golden, CO 80401
USA

p: +1-303-234-9270
f: +1-303-202-6350
Mike.Bode-at-ResAltaTech.com
www.ResAltaTech.com

==============================Original Headers==============================
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Does anyone have experience with the Tucsen 9 MP microscope camera? Is
the computer interface easy to use? Is it easy to change image
resolution back and fourth for focusing and image capture? Is the
measuring software adequate? Can you set the white balance manually and
have it remain constant? How about overall image and video quality? If
you know of any issues with this camera, please let me know on or off list.

==============================Original Headers==============================
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***************************************************************************************
Forwarded from "Ask a Microscopist"
Please remember that the person asking the question is likely
not a member the listserver, and
**any reply should go directly to the poster**
as well as to the list.
Using the "reply" function in your email does *not* send your answer
to the person asking the question.
Please copy their email address from their question.
****************************************************************************************
} Date: Mon, 30 Jan 2012 10:43:44 -0800
} From: {AssociationManagement-at-microscopy.org}
} Reply-To: Jaimie Graham {jaimie_graham-at-cjuhsd.net}
} Subject: Ask-A-Microscopist
}
} Below is the result of your form, submitted on Monday, January 30,
} 2012 at 10:43:40 AM.
}
} realname - Jaimie Graham
} Email - jaimie_graham-at-cjuhsd.net
} ORGANIZATION - Ontario High School
} EDUCATION - 9-12th Grade High School
} LOCATION - Ontario High School
} SUBJECT_OF_QUESTION - SEM
} QUESTION - We have recently received a donated functioning Cambridge
} Instrument Stereoscan 360 SEM. I have used an SEM in college, so
} being one of the only teachers here on campus with any experience I
} have been given the task of figuring out how to put this microscope
} together. I was shipped to us in three main pieces and is currently
} wrapped in plastic. Most of it put together. Will I be able to
} assemble the microscope or do I need to find a company to put it
} together? Is it like a computer where it is simple to figure out
} what plugs into what or am I in way over my head??? Help!
}

==============================Original Headers==============================
1, 24 -- From oshel1pe-at-cmich.edu Mon Jan 30 13:11:28 2012
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Re: donated Cambridge Instruments StereoScan 360 SEM: this looks like a good instrument and it should be nice to have in School, but it's not as easy as a computer to fit together. I would recommend getting professionals to put it together and switch it on. You are going to have to get a Safety certificate before you can use it in class, e.g. an SEM like this does produce X-rays.

The three parts are probably: (1) the electron optical column (heavy metal thing with suspension and a vacuum pumping system), (2) the control electronics console, with monitors and lots of cables around the back and (3) a crate with e.g. high voltage cable, mechanical pump(s), UPS battery system, X-ray detector, computer and accessories.

Try to find manuals, circuit diagrams and ideally installation/performance data; next look into a service company - try searching online. In the best case and with luck you might find someone who can fix things and get the SEM working for around $10K (about a weeks work).

Good luck
--
Robert Keyse
EM Facility
Lehigh University

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Greetings Fellow Microscopists,

The Hooke College of Applied Sciences, located in Westmont, IL, is offering its scanning electron microscopy short course March 26-30, 2012.� In addition to lectures, this course emphasizes hands-on training using state-of-the-art equipment.� For further SEM training details and registration information, please follow the link below:

http://www.hookecollege.com/courses/course.asp?COURSE_ID=42

Best regards-
__________________________________________________
Chris Gorman
Admissions Specialist
Hooke College of Applied Sciences
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7412(fax)
www.hookecollege.com

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Dear Listserv,

I have a chitin hydrogel that I would like to image in TEM. I did some
research and there are reports that standard EM heavy metal stains (osmium,
UA, lead citrate) will stain chitin, however, there are also papers that
indicate that chitin doesn't pick up these stains. I was wondering if
anyone has any experiance staining chitin for TEM.

Ideally, a protocol to stain sections on a grid would be best as I have
some un-stained sections (there is iron oxide in the gel which I wanted
to visualize without any stain). If this isn't possible then I can process
some additional samples.

Thanks very much,
Lyle

--
Lyle Gordon
Department of Materials Science and Engineering
Northwestern University

2220 Campus Drive
Evanston, IL 60208

Tel: (847) 491-3584
lgordon-at-u.northwestern.edu

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7, 23 -- Message-ID: {CAFhTNAmT=KfvvLPP42OZkZTXCe07vLuqsz5PkkP_TsrgRZvOFA-at-mail.gmail.com}
7, 23 -- Subject: Staining Chitin for TEM
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Hello all,
I have following trouble. When starting our Philips CM12 after this night
short blackout I've noticed a strange compressed air noise somewhere around
the diffusion pump. I cannot find out which valve is responsible for it, because
OPCON console cannot be switched to the Vacuum page (pumps and valves scheme)
at this time due to a line scan calibration process.
Thanking you in advance for any suggestion.

Best regards Oldrich

==============================Original Headers==============================
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Dear Oldrich,

I would guess that the problem is coming from the o-rings on the fill and
drain caps of the diffusion pump. These hardened with time and the high
temperatures that they work under. They take on a triangular
cross-section. When the pump cools down, they shrink a fraction, then when
re-heated, because they are no longer elastic, they don't seal fully. The
o-rings will have quite a short life if you use Viton ones- you can buy
black perlast ones for 3x the price of Viton, or Kalrez ones for 4x the
price that have better resistance to oil and heat.

Ben

On 01/02/2012 08:34, "benada-at-biomed.cas.cz" {benada-at-biomed.cas.cz} wrote:

} --------------------------------------------------------------------------
}
} Hello all,
} I have following trouble. When starting our Philips CM12 after this night
} short blackout I've noticed a strange compressed air noise somewhere
} around
} the diffusion pump. I cannot find out which valve is responsible for it,
} because
} OPCON console cannot be switched to the Vacuum page (pumps and valves
} scheme)
} at this time due to a line scan calibration process.
} Thanking you in advance for any suggestion.
}
} Best regards Oldrich

==============================Original Headers==============================
10, 30 -- From ben.micklem-at-pharm.ox.ac.uk Wed Feb 1 04:05:51 2012
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Dear All, our lab is trying to set up some facilities to prepare bio samples for TEM characterization.However, i have little knowledge on bio sample preparation for TEM. On the list of some equipment purchase, we are getting a centrifuge and a rotator. i would like to ask for advice

1) Is it critical to get a centrifuge with a swing out attachment for the centrifuge tube or a fix inclined angle type would be good enough?
2) I realise that the rotator have different degrees e.g 35, 45 and 55 degree, what is the use of different angle on these rotator?

Cheers,
Yee Yan
Nanyang Technological University
FACTS LAB

==============================Original Headers==============================
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4, 35 -- From: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
4, 35 -- Reply-To: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
4, 35 -- Subject: Enquires on Purchasing Centrifuge and Rotator
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Dear Yee Yan,

You bring up an interesting question that broadly affects us all, is it possible for a materials lab to successfully cater to biologists?

I see many universities in the US that set up a materials-dedicated EM lab and then invite biologists to use the equipment. While some of the machines on offer are top of the range microscopes, do they adequately serve the needs of the biologist?

Are there biologists on this server who use materials facilities and are successful on getting all their needs addressed? Do all materials labs have ultramicrotomes available, are there cryosectioning, high pressure freezing and freeze substitution machines available? Can 70 nm thin sections be successfully imaged in a TEM operating at 200kVa? Would the people running the facility be willing to turn down the expensive 200kVa machine to 80k?a for biologists to use?

I think the problem goes further than wondering if a centrifuge will be needed for a biologists - of course it will. Similarly with a rotator. A biologist would find this helpful when resin embedding the cell pellets they formed in the centrifuge. But where is the microwave processor for assisting with specimen embedding too?

Personally I think that biological EM will not survive if shared facilities do not provide expert support to biologists, with only limited knowledge of specimen preparation. There is more to what we do than what expensive microscopes provide.

Some facilities are very successful at catering for materials and biological sciences, but others just can't understand why biologists don't come flooding in. The biological scientists are there, but many of them do not know the best way to get results using the EM. These people need to be helped with advice and instruction, and they are also not usually on this listserver.

My advice would be to find a way to get someone on your staff who has expertise in biological EM and use them for the sort of advice you are looking for. At the very least, invite a biologist into your lab to offer advice on a consultative basis.

Best Regards,

Paul Webster
House Research Institute
2100 W 3rd St
Los Angeles CA 90057
USA

PS
Don't forget to register for our Immuno-Gold-Counting Course taking place in April.

-----Original Message-----
X-from: one_twinklestar-at-yahoo.com.sg [mailto:one_twinklestar-at-yahoo.com.sg]
Sent: Wed 2/1/2012 9:04 AM
To: Webster, Paul

Dear All, our lab is trying to set up some facilities to prepare bio samples for TEM characterization.However, i have little knowledge on bio sample preparation for TEM. On the list of some equipment purchase, we are getting a centrifuge and a rotator. i would like to ask for advice

1) Is it critical to get a centrifuge with a swing out attachment for the centrifuge tube or a fix inclined angle type would be good enough?
2) I realise that the rotator have different degrees e.g 35, 45 and 55 degree, what is the use of different angle on these rotator?

Cheers,
Yee Yan
Nanyang Technological University
FACTS LAB

==============================Original Headers==============================
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Dear Yee Yan,

since I have googled your facility I now know that the primary target of
your Lab-Org will be Materials (http://facts.ntu.edu.sg/about.html ).
I would like to congratulate for the interesting facility map which also
is displayed at (http://facts.ntu.edu.sg/about_lablayout.html)

Seconding Paul Webster's recent reply (and underline the statements he
made) I personally feel that you
perhaps have to (then should) add a biological specimen preparation room
too... if you would like to attract Biologists to use your facility
(nevertheless a compliment for all those machines listed in the map...)

Regarding the secondary questions you were asking, I would like to answer
very briefly:
Centrifuge(s) (for biological preps preferably equipped with a cooling
device) with a "swing out rotor/rotator": would be necessary and IMHO
indispensable e.g. for blood centrifugations (esp. for "Buffy Coat"-Preps).
All spec.-prep. methods for pelleting cellular biological material
(e.g. blood, cell cultures, urinary sediments) will result in a homo-
genous pellet/sediment which can be asservated, conserved and finally
fixed with respective (chem..)fixative(s) and processed that way into
resin.
So called "angle-type" centrifuges might serve too, but produce no even,
orthogonally layered pellets (especially with regard to the even
distribution of "layers" if achievement of such are considered important).

Concerning the different types of "angles":
This means that the higher degree of angle the angle velocity and
thus the translational forces increase (at the end of the test tube)
due to the longer radius of the rotor. So it might well be you'll be
able to achieve higher g-forces on your specs with a 55� than with a 35�
rotor.
Hope this will be of any help with your "decision making"

best regards and greetings to Singapore,

Wolfgang MUSS PhD
SALZBURG, AUSTRIA

} -----Urspr�ngliche Nachricht-----
} Von: one_twinklestar-at-yahoo.com.sg [mailto:one_twinklestar-at-yahoo.com.sg]
} Gesendet: Mittwoch, 01. Februar 2012 18:04
} An: Mu� Wolfgang
} Betreff: [Microscopy] Enquires on Purchasing Centrifuge and Rotator
} -----------------------------------------------------------------------
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}
} Dear All, our lab is trying to set up some facilities to prepare bio
} samples for TEM characterization.
} However, i have little knowledge on bio sample preparation for TEM.
}
} On the list of some equipment purchase, we are getting a centrifuge and
} a rotator.
} i would like to ask for advice
}
} 1) Is it critical to get a centrifuge with a swing out attachment for
} the centrifuge tube or a fix inclined angle type would be good enough?
} 2) I realise that the rotator have different degrees e.g 35, 45 and 55
} degree, what is the use of different angle on these rotator?
}
} Cheers,
} Yee Yan
} Nanyang Technological University
} FACTS LAB
}
}
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10, 39 -- From W.Muss-at-salk.at Wed Feb 1 12:16:54 2012
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Hi Amanda,
First I should disclose that I develop products at Dune Sciences, a
commercial supplier of functionalized TEM grids for advanced applications.
In response to your question restated below, functionalized Carbon grids
will improve and simplify the sample preparation of liposomes. The
positive charge on the grid disperses the liposomes during deposition and
plasma treating is not required as the grids are hydrophilic. You can learn
how functionalization chemistry works by going to the following link.
http://www.dunesciences.com/grids_CSMART.php

SUBJECT_OF_QUESTION - Uranyl Acetate Stain QUESTION - I am staining liposome
with 2 % UA on plasma treated Carbon Type-B grids. Is there anything I can
add to my UA or grid to reduce aggregation ? Thanks.

Thanks,

John Miller
Dune Sciences, Inc.
541-359-1959
jmiller-at-dunesciences.com

-----Original Message-----
X-from: oshel1pe-at-cmich.edu [mailto:oshel1pe-at-cmich.edu]
Sent: Wednesday, February 01, 2012 7:41 AM
To: jmiller-at-dunesciences.com

****************************************************************************
***********
Forwarded from "Ask a Microscopist"
Please remember that the person asking the question is likely not a member
the listserver, and **any reply should go directly to the poster** as well
as to the list.
Using the "reply" function in your email does *not* send your answer to the
person asking the question.
Please copy their email address from their question.
****************************************************************************
************

realname - Amanda Lever
Email - {mailto:lever.amanda1-at-gmail.com} lever.amanda1-at-gmail.com
ORGANIZATION - Draper Laboratory
EDUCATION - Graduate College
LOCATION - Boston, MA
SUBJECT_OF_QUESTION - Uranyl Acetate Stain QUESTION - I am staining liposome
with 2 % UA on plasma treated Carbon Type-B grids. Is there anything I can
add to my UA or grid to reduce aggregation ? Thanks.

==============================Original Headers==============================
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We have a 120kV FEI Tecnai 12 Twin TEM which was chosen to serve multiple purpose projects. On installation the scope was rated at 2nm. We have stereo-imaged 500nm sections, 70nm sections of epoxy-embedded silica gel impregnated with Cobalt oxide deposits, and nano-particulates on C-coated grids. For a relatively non-analytic, vanilla, 120kV instrument, I believe that is not bad. I routinely view thin sections (60-80nm) of biological tissue post-fixed with osmium and permit the 1k Gatan to add the contrast which is usually quite satisfactory. The vacuum is usually in the middle of 10-8 T which is also more than satisfactory. We did not spec a 'bio' lens configuration, because we wanted to be flexible.
We have a lab equipped as can be seen on our web site.

Most of our users are not biologists, but we can certainly do biologic specimens without any problem. The only difference I have imposed on the use of the TEM is that all biological specimens and those nano particles produced by wet chemistry and not presented in light alcohols are used with the LN2 coldfinger to prevent general contamination in the stage area.

Short of the accessories for vitreous quenching, the biggest difference will lie with the biological core facility that cannot be used by materials scientists - no matter what the size/expense.

Finally, having said all of the above, there are choices in instruments, because there is variety of need. One can stretch one's flexibility so far that it excludes the need one was supposed to address in the first place. A core facility with a Titan and a 120kV Bio-Twin would serve both material and biological investigators. Under what conditions would a biologist need wave-length resolution and not, in truth, be working on bio-materials.

As an example, if I had the money and I wanted to purchase a Titan with 10 years of service, I might spend it and have the package delivered to someone who has a discernible need. I, even when I try with diligence, cannot uncover a need/use for myself. So, I would not recommend such an acquisition, even if it were offered, if I could not worry up a need to have it. Then, of course, the ceiling here is not nearly high enough, and even though on bedrock, and lacking subways, there are still trucks and busses, AND, where I now need only two rooms for the TEM, I would need at least three for a Titan, and perhaps for a 200kV with no observation chamber as well.

The Dean of a college has a materials-oriented core facility for which s/he is responsible (services, maintenance, and cost recovery). S/he charges the Deans of other colleges whose Faculty use the facility. The President owns the Colleges and the cores, and spends money on an accounting scheme which issues bills for use to Departments and Individuals who have external funding. There is no mechanism to charge for undergraduate courses, which, thus, do not usually fit into such a scheme. So the core is limited to research use, and many bright students choose business and finance.

Cheers,

Fred Monson

P.S. The day is almost over, and I look forward to the continuing mechanical and optical attention I am paying to one of my antique Leitz 'Research' Microscopes. I could NEVER do that with a Titan, nor even with our Tecnai. How sad.... I, not a Tinker, Tailor, nor a Soldier be, but merely a mechanic - just like me Dad. God bless 'im!

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

If one looks long enough, one may actually see something frozen in time that flashes one's mind to a new state.
1. The day that I asked myself how the hydrogen atom's mass could actually be: 1.0008.
2. The day, while looking at a slide of some tissue, I saw a leukocyte that had been caught sneaking in or out of a capillary: http://www.annualreviews.org/doi/abs/10.1146/annurev-pathol-011110-130224

-----Original Message-----
X-from: PWebster-at-hei.org [mailto:PWebster-at-hei.org]
Sent: Wednesday, February 01, 2012 12:36 PM
To: Monson, Frederick

Dear Yee Yan,

You bring up an interesting question that broadly affects us all, is it possible for a materials lab to successfully cater to biologists?

I see many universities in the US that set up a materials-dedicated EM lab and then invite biologists to use the equipment. While some of the machines on offer are top of the range microscopes, do they adequately serve the needs of the biologist?

Are there biologists on this server who use materials facilities and are successful on getting all their needs addressed? Do all materials labs have ultramicrotomes available, are there cryosectioning, high pressure freezing and freeze substitution machines available? Can 70 nm thin sections be successfully imaged in a TEM operating at 200kVa? Would the people running the facility be willing to turn down the expensive 200kVa machine to 80k?a for biologists to use?

I think the problem goes further than wondering if a centrifuge will be needed for a biologists - of course it will. Similarly with a rotator. A biologist would find this helpful when resin embedding the cell pellets they formed in the centrifuge. But where is the microwave processor for assisting with specimen embedding too?

Personally I think that biological EM will not survive if shared facilities do not provide expert support to biologists, with only limited knowledge of specimen preparation. There is more to what we do than what expensive microscopes provide.

Some facilities are very successful at catering for materials and biological sciences, but others just can't understand why biologists don't come flooding in. The biological scientists are there, but many of them do not know the best way to get results using the EM. These people need to be helped with advice and instruction, and they are also not usually on this listserver.

My advice would be to find a way to get someone on your staff who has expertise in biological EM and use them for the sort of advice you are looking for. At the very least, invite a biologist into your lab to offer advice on a consultative basis.

Best Regards,

Paul Webster
House Research Institute
2100 W 3rd St
Los Angeles CA 90057
USA

PS
Don't forget to register for our Immuno-Gold-Counting Course taking place in April.

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Dear All, our lab is trying to set up some facilities to prepare bio samples for TEM characterization.However, i have little knowledge on bio sample preparation for TEM. On the list of some equipment purchase, we are getting a centrifuge and a rotator. i would like to ask for advice

1) Is it critical to get a centrifuge with a swing out attachment for the centrifuge tube or a fix inclined angle type would be good enough?
2) I realise that the rotator have different degrees e.g 35, 45 and 55 degree, what is the use of different angle on these rotator?

Cheers,
Yee Yan
Nanyang Technological University
FACTS LAB

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Hi Erin,

I use LR white almost exclusively for immunolabeling in plant tissue and have very good luck with it. The protocol that I use I got from Kevin Vaughn, so I would suggest that you check out a few of his papers. Some of his earlier work was on immunolabeling various proteins in chloroplasts, so they may be especially relevant to you, though the later papers may have slightly more up-to-date protocols.

If you have any trouble finding his papers or need more detail on any of the steps of the protocol, just let me know.

Andy

_____________________
Andrew J Bowling, PhD
Discovery Research
Dow AgroSciences
9330 Zionsville Rd
Indianapolis, IN 46268
317-337-3878

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Email: stempies-at-muohio.edu Name: Erin Stempinski

Organization: Miami University

Title-Subject: [Filtered] Resins for immunocytochemistry

Message: Dear Listserv,

I am looking at immunogold labeling proteins in Arabidopsis chloroplasts for TEM and was wondering
if anyone on the Listserv had experience with the differences in immunocytochemistry resins. I'm
interested in knowing how well Unicryl, LR White, and LR Gold have performed in other people's
experience with immunogold labeling.

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Hi Lyle and all,
Try the protocol in Lingle, W.L. (1989). Enhanced Staining of the
Basidiomycete Panellus stypticus Prepared for TEM by Freeze
Substitution. Cryotogamic Botany 1, 236-242. Wilma did a modification
of the Thiery stain and this method stains chitin nicely. I couldn't
find the journal article on-line but I can snail mail a copy to you if
you'd like it.
best,
Beth

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Just a thought - the fluorescent lectins are very good at binding
specifically to chitin, e.g. fluorescent wheat germ agglutinin. Would a
gold-tagged one work in TEM? I don't know if they exist, but it would be
very straightforward to do the staining.
cheers,
Rosemary

Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
M 61 2 420 972 028
E rosemary.white-at-csiro.au

On 2/02/12 8:40 AM, "beth-at-plantbio.uga.edu" {beth-at-plantbio.uga.edu} wrote:

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10, 31 -- Subject: Re: [Microscopy] staining chitin
10, 31 -- From: Rosemary White {rosemary.white-at-csiro.au}
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Hi Rosemary,

Good idea with the lectins. However, I would not use lectins directly
coupled to gold particles. Instead, for EM labeling I would probably apply
unbound lectin to the sections and then specific anti-lectin antibodies
followed by the gold probe. Then there would be no concern about the gold
particles disassociating from the lectin during storage.

Of course, if going the antibody route, why not use anti-chitin antibodies?

Regards

Paul.

Paul Webster, Ph.D.
House Research Institute
2100 West Third Street
Los Angeles
CA 90057
(213) 273 8026

On 2/1/12 2:13 PM, "rosemary.white-at-csiro.au" {rosemary.white-at-csiro.au}
wrote:

} microscopy-at-microscopy.com

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It's Friday and if the prospect of a week-end isn't enough, Wired has come through. No, they're not all photographs. The article is about Science Visualizations and some great images. I'm disappointed there's no fusion preps in PLM but the cucumber skin barbs make up for it.
Here's the link.

http://www.wired.com/wiredscience/2012/02/science-visualizations-2011/?utm_source=feedburner&utm_medium=feed&utm_campaign=Feed%3A+wired%2Findex+%28Wired%3A+Index+3+%28Top+Stories+2%29%29&pid=3031

One can't help wonder what the folks at Wired think of the sudden influx of web traffic for specific article.

Frank Karl
Microscopist
ARDL
2887 Gilchrist Road
Akron, Ohio 44305
330-794-6600

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Hi Patricia,

I posed the same question to my Oxford service engineer and he told me
that they recommend leaving the detector in operate mode. I haven't
been able to detect any adverse effects in detector performance.

Bryan

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} Title-Subject: [Filtered] SDD - standby or operate mode
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} Message: We have a new SDD Oxford detector on our S-4700. I was
} wondering if most owners of these systems keep them in Operate mode, or
} only cool them down when users are scheduled.
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--
_________________________________________________________________
Bryan R. Bandli

Research Instrumentation Laboratory Manager
University of Minnesota, Duluth
229 Heller Hall
1114 Kirby Dr.
Duluth, MN� 55812
218-726-7362
==================================================================

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Hi Patricia,

We also have an Oxford SDD and the technologist who installed it
recommended that we keep it in standby mode when not being used in the
near future.

Translation: if it would be used within 12-24 hr, keep it chilled. Any
longer periods, go to standby.

My personal opinion is keeping it chilled all the time is putting an
unnecessary "strain" on the Peltier electronics.

--
John J. Bozzola, Ph.D.
Professor & Blissfully Retired Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

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Patricia

I have spoken with Oxford's applications people and they suggest leaving it
in Standby except when needed. It takes no more than 5 minutes to cool down
to the operating temperature.

Alan

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Alan W Nicholls, PhD
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In recent years we have been getting requests to image functionalized
nanoparticles, such as 10 nm gold with proteins or peptides attached.
Generally the clients want to know how thick or large the organic
"shell" is, or how the protein/peptide is arranged on the surface.
Most of our experience has been with either inorganic particles (which
we view without staining) or larger organic structures such as viral
procapsids or large proteins (e.g., myosin) or protein assemblies,
which we negative stain. My questions for this list are as follows:

- have others on the list had good luck imaging these kinds of
conjugates at the EM level, and if so, using what techniques?
- how much organic material must one have associated with the
nanoparticle to see it by negative staining or other methods? For
example, how hard is it to detect a 150 kD protein on a 10 nm gold
particle? An example would be IgG attached to a 10 nm gold particle.
- can you suggest any publications or review articles on the subject?
For some reason I'm not having much luck with my Pubmed search
strategies. A recent paper by Cao and Mao seems to deal mainly with
small gold particles on much larger protein aggregates, but perhaps
there are others out there.

Many thanks (in advance) for your suggestions.

Dr. Marie E. Cantino
Associate Professor of Physiology and Neurobiology
Director, Electron Microscopy Laboratory
University of Connecticut, Unit 3242
Phone: 860-486-3588
Fax: 860-486-6369

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Group,

Would like to explore the experiences and words of wisdom on the idea of having a "core analytical facility " on a University campus.
Questions I am exploring are:
How is this idea presented to administrators as a needed addition to support research activities across many disciplines? Or is it?
How are such facilities managed and what are reasonable staffing levels.
What drives the tool set? Is it internal existing research or potential outside research partnerships?
Finally most important is addressing how to pay for and continually support such a facility?

Thanks for any thoughts or ideas on the subject.

Roy Beavers
Southern Methodist University
Department of Earth Sciences
3225 Daniel Ave
Dallas, TX� 75205
Voice: 214-768-2756
Email: rbeavers-at-smu.edu

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Lyle:

I have done a lot of work with Fungal samples whose walls are mostly
chitin. The walls routinely stained well with OsO4, Uac, and PbCit - sorry I
never did a test to see which predominantly stained the chitin. However, I
have used Barium Permanganate specifically for stainging fungal walls for
TEM and it does a great job. Original reference is:

Hoch, H. C. 1977. Use of permanganate of increase the electron opacity
of fungal walls. Mycologia 69:1209-2113.

On 31 Jan 2012 at 15:34, lgordon-at-gmail.com wrote:

}
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} Dear Listserv,
}
} I have a chitin hydrogel that I would like to image in TEM. I did some
} research and there are reports that standard EM heavy metal stains
} (osmium, UA, lead citrate) will stain chitin, however, there are also
} papers that indicate that chitin doesn't pick up these stains. I was
} wondering if anyone has any experiance staining chitin for TEM.
}
} Ideally, a protocol to stain sections on a grid would be best as I
} have some un-stained sections (there is iron oxide in the gel which I
} wanted to visualize without any stain). If this isn't possible then I
} can process some additional samples.
}
} Thanks very much,
} Lyle
}
} --
} Lyle Gordon
} Department of Materials Science and Engineering
} Northwestern University
}
} 2220 Campus Drive
} Evanston, IL 60208
}
} Tel: (847) 491-3584
} lgordon-at-u.northwestern.edu
}
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Richard E. Edelmann, Ph.D., Director
Center for Advanced Microscopy & Imaging
9C Upham Hall
Miami University, Oxford, OH 45056
Ph: 513.529.5712 Fax: 513.529.4243
E-mail: edelmare-at-muohio.edu
http://www.cami.muohio.edu

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Dear Colleagues,

On behalf of the Microanalysis Society (MAS), I invite you to the 3rd Topical Conference on EBSD, which will be held at Carnegie Mellon University on June 19-21, 2012. We have fantastic new speakers, and we are once again providing a comprehensive tutorial on EBSD that will include live demonstrations. For more information, and to register, please visit our website:

http://www.microbeamanalysis.org/topical-conferences/ebsd-2012/

Space is limited, so please register early. We look forward to seeing you in Pittsburgh!

Best regards,
Andrew Deal

MAS Director,
EBSD 2012 Organizing Committee Chair

Thanks,
Andy
********************************************
Andrew Deal, PhD
Materials Scientist
GE Global Research
One Research Circle
Building K1, Room 1D-37A
Niskayuna, NY 12309
US
T +1 518 387 5456
F +1 518 387 6905
********************************************

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Hi All,

For any of you out there who think they might like to try administration, a
position has opened up at Purdue and we (other interested microscopists on
campus) are hoping for someone who is a strong advocate of microscopy.

Take a look and pass this on if you know of someone who might fit the bill.

++++++++++++++++++++++++++++++++++++++++++++++

Head and Professor
School of Materials Engineering

Purdue University is seeking nominations and applications for the position
of Head of the School of Materials Engineering. A dynamic leader is sought
to advance the School�s nationally-ranked program and to enhance its
international and national impact through a continuing commitment to
excellence in discovery, learning and engagement. The Head provides vision
and leadership to the faculty, students, staff, alumni, and other
stakeholders of the School, and will have the opportunity to shape and
implement the strategic plan for the School and the College of Engineering.

The Head must have a Ph.D. in Materials Science and Engineering or related
disciplines, qualify for an appointment at the full professor level with
tenure, have a distinguished academic, government or industrial record, and
demonstrate strong leadership and collaborative skills. Candidates should
have a clear understanding of the current needs and future direction of the
materials science and engineering profession, possess a commitment to
diversity and collaboration, and be skilled in administration, student
relations, mentoring, and alumni development.

The faculty members in the School of Materials Engineering are located in
the Neil Armstrong Hall of Engineering and the Birck Nanotechnology Center,
which provide outstanding facilities for teaching and research. The School
currently has 116 undergraduate students, 89 graduate students and postdocs,
and 20 faculty members. The School�s faculty members have strong connections
to research centers in Purdue�s Discovery Park, as well as with the Colleges
of Agriculture, Liberal Arts, Pharmacy, Management, and Science.

The College of Engineering at Purdue consists of over 7300 undergraduate
students, nearly 2,900 graduate students, and 359 faculty members. Purdue is
one of the nation�s leading land-grant universities with a full range of
academic majors and an enrollment of over 40,000 students on the West
Lafayette campus.

An application should include: (1) a three-page personal statement
addressing the applicant's vision, administrative philosophy, experience and
qualifications; (2) a curriculum vitae; and (3) names and contact
information for at least three references. Applications and inquiries will
be kept confidential. Applicants will be notified before references are
contacted. Electronic submission is preferred and should be uploaded at
https://engineering.purdue.edu/Engr/InfoFor/Employment. Screening will
commence February 1, 2012, and continue until this position is filled.
Nominations and questions regarding the position can be addressed to Chair,
Materials Engineering Head Search Committee, Purdue University, Neil
Armstrong Hall of Engineering, 701 W. Stadium Avenue, West Lafayette, IN
47907-2045, Phone: 765-494-5012, Email: mse-search-at-ecn.purdue.edu. A
background check will be required for employment in this position. Purdue
University is an equal opportunity/equal access/affirmative action employer
fully committed to achieving a diverse workforce.

Debby

--
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy/

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Hi all,
We have an otherwise perfectly fine Sorvall RC2-B which failed it's recent safety check because the inner plastic cover of the lid has a crack. Of course, no parts can be found for that centrifuge anymore.
Thus, if you are trashing your old RC2-B, it would be great if we could get the plastic cover which is just screwed to the lid from the inside.

Thanks in advance for any help,

Anja Seybert
Research Scientist
Frankfurt am Main, Germany

==============================Original Headers==============================
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Hi Mary!
�
I am afraid I am not a specialist of this technique but I know how it works and I would like to just make some general comments.
Others with more experience may want to give a more precise answer.
�
This method involves statistical analysis, which means the more you sample, the more precise your number.
In other words, the precision of your analysis will basically depend on how much time you want to put into your counting (how many grids/fields you actually count).
Based on that, I assume that you may be able to count fewer numbers (decrease your lower limit) by counting many grids.
�
Best regards,
Stephane
�
�
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Email: mary.walker-at-csl.com.au Name: mary walker

Organization: csl limited

Title-Subject: [Filtered] counting virus particles

Message: Do any listers have experience with counting virus particles
with latex beads?
Is it possible to do better than order of magnitude?
What is the minimum number of particles per ml that can be counted?
What is the lower limit?

thanks in advance

� Login Host: 203.10.36.68
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Hi Marie,

I like these projects which put the techniques to their limits.
I think the reason why you have difficulties finding publications on this matter and that the question is not trivial at all.
First of all, and to start with a relaxed note, the mixing of kDa und nm in one sentence made me smile because it makes as much sense as asking if 1kg of something fits in one meter.
The Dalton is a unit of mass and does not give a good indication of the dimension or the shape of a molecule.
That said, I seem to remember from the past answers of our emminent immunoglobulinologist Jan Leunissen that the size of an IgG is several nm, I think more than 5nm.
Which means, in your case, that you should not expect more that a few molecules per nanoparticle, except if they form multilayers.
As for the techniques, I wonder if a SEM could resolve this structure. Although I have experience with analytical low res SEMs, I don't know the limits of modern FEG SEMs.
I would be happy if some listers would care to share�their opinion on this matter.
It seems that AFM (atomic force microscopy) would be able to resolve this structure, but I wonder if it would be possible in this condition because AFM works on flat substrate and here you have round particles.
It would probably be possible to find some answers using cry-microscopy, the same way scientists use it to resolve viral structures at several anstrom resolution. It probably depends on how much time and money your clients are wanting to spend and how critical this information is for them.

Regards,
Stephane
�
----- Original Message -----
X-from: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
To: nizets2-at-yahoo.com
Cc:
Sent: Friday, February 3, 2012 8:10 PM

In recent years we have been getting requests to image functionalized�
nanoparticles, such as 10 nm gold with proteins or peptides attached.�
Generally the clients want to know how thick or large the organic�
"shell" is, or how the protein/peptide is arranged on the surface.�
Most of our experience has been with either inorganic particles (which�
we view without staining) or larger organic structures such as viral�
procapsids or large proteins (e.g., myosin) or protein assemblies,�
which we negative stain.� My questions for this list are as follows:

- have others on the list had good luck imaging these kinds of�
conjugates at the EM level, and if so, using what techniques?
- how much organic material must one have associated with the�
nanoparticle to see it by negative staining or other methods?� For�
example, how hard is it to detect a 150 kD protein on a 10 nm gold�
particle?� An example would be IgG attached to a 10 nm gold particle.
- can you suggest any publications or review articles on the subject?�
For some reason I'm not having much luck with my Pubmed search�
strategies.� A recent paper by Cao and Mao seems to deal mainly with�
small gold particles on much larger protein aggregates, but perhaps�
there are others out there.

Many thanks (in advance) for your suggestions.

Dr. Marie E. Cantino
Associate Professor of Physiology and Neurobiology
Director, Electron Microscopy Laboratory
University of Connecticut, Unit 3242
Phone: 860-486-3588
Fax: 860-486-6369

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Hi Ravi,

Movement of the image when changing the objective lens (i.e. FOCUS) is
an indication that the beam is going through the objective lens at an
angle, i.e. not on the optic axis. The correction step is usually
referred to as a "Rotation Calibration" or Objective lens "Current
Centering".

It is difficult to describe the exact steps to correct this without
knowing which microscope you have since the knobs will vary from
microscope to microscope.

Many microscopes have a direct alignment step labelled "rotation center"
which oscillates the objective lens. Start with the image close to
focus, engage the "rotation center" function (start the objective lens
current oscillation) and adjust the correction knobs to minimize the
image movement.

For a non-computerized microscope, there may be a set of knobs labelled
"rotation center". In this case, focus the image and set a recognizable
feature in the center. Then defocus and watch the feature shift away
from the center. Use the "rotation center" knobs to recenter the
feature while the image is defocused. Repeat if necessary.

Good luck,
Henk

At 2/6/2012 9:11 AM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Email: ravi.thakkar369-at-gmail.com Name: Ravindra Thakkar
}
} Title-Subject: [Filtered] TEM : Focusing Problem.
}
} Message: Image is not getting focused. The image is drastically moving
} while focusing, it moving highly even at low focus step& at low
} magnification.
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} So how to resolve this problem.
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--
Hendrik O. Colijn
www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
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"Time is that quality of nature which keeps things from happening all at
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Hi Stefan,

I do understand the difference between mass and length.

The problem is that the clients usually can only give us the molecular
weight and (if we're lucky) the number of molecules per particle. If
they knew how the proteins/peptides were arranged or the linear
dimensions of the coating they wouldn't need to come to me. What I'm
trying to find out from the listserver or the literature is whether we
can reasonably expect to be able to detect a medium sized protein such
as IgG on an electron dense gold particle, given the various artifacts
of techniques such as negative staining.

We only have TEM (w/a tungsten emitter) and FESEM in our facility, but
I can send them elsewhere for AFM.

Marie

On Feb 6, 2012, at 5:33 AM, nizets2-at-yahoo.com wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi Marie,
}
} I like these projects which put the techniques to their limits.
} I think the reason why you have difficulties finding publications on
} this matter and that the question is not trivial at all.
} First of all, and to start with a relaxed note, the mixing of kDa
} und nm in one sentence made me smile because it makes as much sense
} as asking if 1kg of something fits in one meter.
} The Dalton is a unit of mass and does not give a good indication of
} the dimension or the shape of a molecule.
} That said, I seem to remember from the past answers of our emminent
} immunoglobulinologist Jan Leunissen that the size of an IgG is
} several nm, I think more than 5nm.
} Which means, in your case, that you should not expect more that a
} few molecules per nanoparticle, except if they form multilayers.
} As for the techniques, I wonder if a SEM could resolve this
} structure. Although I have experience with analytical low res SEMs,
} I don't know the limits of modern FEG SEMs.
} I would be happy if some listers would care to share their opinion
} on this matter.
} It seems that AFM (atomic force microscopy) would be able to resolve
} this structure, but I wonder if it would be possible in this
} condition because AFM works on flat substrate and here you have
} round particles.
} It would probably be possible to find some answers using cry-
} microscopy, the same way scientists use it to resolve viral
} structures at several anstrom resolution. It probably depends on how
} much time and money your clients are wanting to spend and how
} critical this information is for them.
}
} Regards,
} Stephane
}
} ----- Original Message -----
} X-from: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
} To: nizets2-at-yahoo.com
} Cc:
} Sent: Friday, February 3, 2012 8:10 PM
} Subject: [Microscopy] TEM of functionalized nanoparticles
}
}
}
}
} ----------------------------------------------------------------------------
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} America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} In recent years we have been getting requests to image functionalized
} nanoparticles, such as 10 nm gold with proteins or peptides attached.
} Generally the clients want to know how thick or large the organic
} "shell" is, or how the protein/peptide is arranged on the surface.
} Most of our experience has been with either inorganic particles (which
} we view without staining) or larger organic structures such as viral
} procapsids or large proteins (e.g., myosin) or protein assemblies,
} which we negative stain. My questions for this list are as follows:
}
} - have others on the list had good luck imaging these kinds of
} conjugates at the EM level, and if so, using what techniques?
} - how much organic material must one have associated with the
} nanoparticle to see it by negative staining or other methods? For
} example, how hard is it to detect a 150 kD protein on a 10 nm gold
} particle? An example would be IgG attached to a 10 nm gold particle.
} - can you suggest any publications or review articles on the subject?
} For some reason I'm not having much luck with my Pubmed search
} strategies. A recent paper by Cao and Mao seems to deal mainly with
} small gold particles on much larger protein aggregates, but perhaps
} there are others out there.
}
} Many thanks (in advance) for your suggestions.
}
} Dr. Marie E. Cantino
} Associate Professor of Physiology and Neurobiology
} Director, Electron Microscopy Laboratory
} University of Connecticut, Unit 3242
} Phone: 860-486-3588
} Fax: 860-486-6369
}
}
} ==============================Original
} Headers==============================
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} 15, 42 -- Date: Mon, 6 Feb 2012 02:28:29 -0800 (PST)
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} 15, 42 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com}
} 15, 42 -- Subject: Re: [Microscopy] TEM of functionalized
} nanoparticles
} 15, 42 -- To: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
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Dr. Marie E. Cantino
Associate Professor of Physiology and Neurobiology
Director, Electron Microscopy Laboratory
University of Connecticut, Unit 3242
Phone: 860-486-3588
Fax: 860-486-6369

==============================Original Headers==============================
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hello Marie,

I am not certain this answers your question but you should be able to detect IgG attached to an electron dense gold particle by negative stain EM. Antibodies are commonly seen attached to virus particles and to colloidal gold particles when negative stain EM is done. Anything you would likely see in your indicated study by AFM can be seen by negative stain EM. All techniques have various artifacts including light-scatter, thin-section EM, AFM, FESEM, and cryo-EM. The key to EM and techniques in general is to learn what is real and what is artifact. It often takes time and work to develop proficiency with many techniques though. Good luck with this , it should be possible.

Charles Humphrey, Ph. D.
CDC
Atlanta, GA 30333

-----Original Message-----
X-from: marie.cantino-at-uconn.edu [mailto:marie.cantino-at-uconn.edu]
Sent: Monday, February 06, 2012 9:49 AM
To: Humphrey, Charles (CDC/OID/NCEZID)

Hi Stefan,

I do understand the difference between mass and length.

The problem is that the clients usually can only give us the molecular weight and (if we're lucky) the number of molecules per particle. If they knew how the proteins/peptides were arranged or the linear dimensions of the coating they wouldn't need to come to me. What I'm trying to find out from the listserver or the literature is whether we can reasonably expect to be able to detect a medium sized protein such as IgG on an electron dense gold particle, given the various artifacts of techniques such as negative staining.

We only have TEM (w/a tungsten emitter) and FESEM in our facility, but I can send them elsewhere for AFM.

Marie

On Feb 6, 2012, at 5:33 AM, nizets2-at-yahoo.com wrote:

}
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} Hi Marie,
}
} I like these projects which put the techniques to their limits.
} I think the reason why you have difficulties finding publications on
} this matter and that the question is not trivial at all.
} First of all, and to start with a relaxed note, the mixing of kDa und
} nm in one sentence made me smile because it makes as much sense as
} asking if 1kg of something fits in one meter.
} The Dalton is a unit of mass and does not give a good indication of
} the dimension or the shape of a molecule.
} That said, I seem to remember from the past answers of our emminent
} immunoglobulinologist Jan Leunissen that the size of an IgG is several
} nm, I think more than 5nm.
} Which means, in your case, that you should not expect more that a few
} molecules per nanoparticle, except if they form multilayers.
} As for the techniques, I wonder if a SEM could resolve this structure.
} Although I have experience with analytical low res SEMs, I don't know
} the limits of modern FEG SEMs.
} I would be happy if some listers would care to share their opinion on
} this matter.
} It seems that AFM (atomic force microscopy) would be able to resolve
} this structure, but I wonder if it would be possible in this condition
} because AFM works on flat substrate and here you have round particles.
} It would probably be possible to find some answers using cry-
} microscopy, the same way scientists use it to resolve viral structures
} at several anstrom resolution. It probably depends on how much time
} and money your clients are wanting to spend and how critical this
} information is for them.
}
} Regards,
} Stephane
}
} ----- Original Message -----
} X-from: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
} To: nizets2-at-yahoo.com
} Cc:
} Sent: Friday, February 3, 2012 8:10 PM
} Subject: [Microscopy] TEM of functionalized nanoparticles
}
}
}
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}
} In recent years we have been getting requests to image functionalized
} nanoparticles, such as 10 nm gold with proteins or peptides attached.
} Generally the clients want to know how thick or large the organic
} "shell" is, or how the protein/peptide is arranged on the surface.
} Most of our experience has been with either inorganic particles (which
} we view without staining) or larger organic structures such as viral
} procapsids or large proteins (e.g., myosin) or protein assemblies,
} which we negative stain. My questions for this list are as follows:
}
} - have others on the list had good luck imaging these kinds of
} conjugates at the EM level, and if so, using what techniques?
} - how much organic material must one have associated with the
} nanoparticle to see it by negative staining or other methods? For
} example, how hard is it to detect a 150 kD protein on a 10 nm gold
} particle? An example would be IgG attached to a 10 nm gold particle.
} - can you suggest any publications or review articles on the subject?
} For some reason I'm not having much luck with my Pubmed search
} strategies. A recent paper by Cao and Mao seems to deal mainly with
} small gold particles on much larger protein aggregates, but perhaps
} there are others out there.
}
} Many thanks (in advance) for your suggestions.
}
} Dr. Marie E. Cantino
} Associate Professor of Physiology and Neurobiology Director, Electron
} Microscopy Laboratory University of Connecticut, Unit 3242
} Phone: 860-486-3588
} Fax: 860-486-6369
}
}
} ==============================Original
} Headers==============================
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Dr. Marie E. Cantino
Associate Professor of Physiology and Neurobiology Director, Electron Microscopy Laboratory University of Connecticut, Unit 3242
Phone: 860-486-3588
Fax: 860-486-6369

==============================Original Headers==============================
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Image moving while focusing means that electron beam is not passing
through the optical center of the lens. Either there is a problem with
alignment electronics and it is pushed beam aside from its optimal path,
or there is a problem with electron-optical elements inside of the
column (maybe something got knocked out of alignment, or there is a
large particle that got charged and distorted the field, or something
else of similar nature).

In any case - look like you need to call a serviceman...

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

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}
} Title-Subject: [Filtered] TEM : Focusing Problem.
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One thing to look for in the electronics readout system (perhaps a
computer with memory) is the alignment numbers, giving gun alignment
numbers (two sets for X and two sets of Y) and the condenser alignment
(likewise) and perhaps Projector alignment, even image shift on some
models. Basically you would love to have a set of "known good" numbers
for reference. By comparing the last known good alignment numbers with
the present numbers you can eliminate the possibility of the previous
user being rather too keen to perform 'dark field' experiments.

As previous posters mention it is the condenser lens (CL) TILT alignment
that might be in error, another possibility is that the sample is bent
horribly and the lens is focusing at a value that is nowhere near it's
optimal value (have you checked the eucentric height?). Again having
values of lens currents recorded after a good session can be useful and
many service people do just that, by checking the performance report
from the installation.

Is the Gun tilt (filament image), condenser aperture centering, and spot
size adjustments all perfectly normal?

Good luck,
Rob Keyse

On 2/6/2012 10:53 AM, vray-at-partbeamsystech.com wrote:
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}
} Image moving while focusing means that electron beam is not passing
} through the optical center of the lens. Either there is a problem with
} alignment electronics and it is pushed beam aside from its optimal path,
} or there is a problem with electron-optical elements inside of the
} column (maybe something got knocked out of alignment, or there is a
} large particle that got charged and distorted the field, or something
} else of similar nature).
}
} In any case - look like you need to call a serviceman...
}
} Valery Ray
} =================================
} PBS&T, MEO Engineering Co., Inc.
} 290 Broadway, Suite 298
} Methuen, MA 01844 USA
} Phone: +1-978-296-5063 - leave a message with call-back number
} US Mobile: +1-978-305-0479
} Skype: pbstmeo
} E-mail: vray-at-partbeamsystech.com
} Web: www.partbeamsystech.com
} Web: www.freudlabs.com
}
} On 2/6/2012 9:08 AM, microscopylistserver-noreply-at-microscopy.com wrote:
} } ----------------------------------------------------------------------------
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} } Email: ravi.thakkar369-at-gmail.com Name: Ravindra Thakkar
} }
} } Title-Subject: [Filtered] TEM : Focusing Problem.
} }
} } Message: Image is not getting focused. The image is drastically moving
} } while focusing, it moving highly even at low focus step& at low
} } magnification.
} }
} } So how to resolve this problem.
} }
} } Login Host: 14.139.97.76
} } ---------------------------------------------------------------------------
} }

--
Robert Keyse
EM Facility
Whitaker Laboratory
5 East Packer Avenue
Bethlehem
PA 18015
USA

Tel. +1 610 758 4283
Fax +1 610 758 4244

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Hello Marie,
I agree with Charles: "you should be able to see IgG .... by negative
staining". However, I have never used negative staining for samples containing
colloidal gold particles bigger than 10 nm.
In my hand (~5 to ~10 nm gold particles on thin carbon support film), 1% or 2%
ammonium molybdate or its mixture with 0.1% trehalose worked better than
classic 2% uranyl acetate. Meniscus of uranyl acetate around the gold particle
was usually too dense to see IgG attached to it at 80 kV and tungsten cathode.

Best regards Oldrich

Prague
Czech Republic

On Monday 06 of February 2012 16:23:06 you wrote:
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} Hello Marie,
}
} I am not certain this answers your question but you should be able to
} detect IgG attached to an electron dense gold particle by negative stain
} EM. Antibodies are commonly seen attached to virus particles and to
} colloidal gold particles when negative stain EM is done. Anything you
} would likely see in your indicated study by AFM can be seen by negative
} stain EM. All techniques have various artifacts including light-scatter,
} thin-section EM, AFM, FESEM, and cryo-EM. The key to EM and techniques in
} general is to learn what is real and what is artifact. It often takes
} time and work to develop proficiency with many techniques though. Good
} luck with this , it should be possible.
}
} Charles Humphrey, Ph. D.
} CDC
} Atlanta, GA 30333
}
} -----Original Message-----
} X-from: marie.cantino-at-uconn.edu [mailto:marie.cantino-at-uconn.edu]
} Sent: Monday, February 06, 2012 9:49 AM
} To: Humphrey, Charles (CDC/OID/NCEZID)
} Subject: [Microscopy] TEM of functionalized nanoparticles
}
}
}
}
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} Hi Stefan,
}
} I do understand the difference between mass and length.
}
} The problem is that the clients usually can only give us the molecular
} weight and (if we're lucky) the number of molecules per particle. If they
} knew how the proteins/peptides were arranged or the linear dimensions of
} the coating they wouldn't need to come to me. What I'm trying to find out
} from the listserver or the literature is whether we can reasonably expect
} to be able to detect a medium sized protein such as IgG on an electron
} dense gold particle, given the various artifacts of techniques such as
} negative staining.
}
} We only have TEM (w/a tungsten emitter) and FESEM in our facility, but I
} can send them elsewhere for AFM.
}
} Marie
}
} On Feb 6, 2012, at 5:33 AM, nizets2-at-yahoo.com wrote:
} } ----------------------------------------------------------------------
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} }
} } Hi Marie,
} }
} } I like these projects which put the techniques to their limits.
} } I think the reason why you have difficulties finding publications on
} } this matter and that the question is not trivial at all.
} } First of all, and to start with a relaxed note, the mixing of kDa und
} } nm in one sentence made me smile because it makes as much sense as
} } asking if 1kg of something fits in one meter.
} } The Dalton is a unit of mass and does not give a good indication of
} } the dimension or the shape of a molecule.
} } That said, I seem to remember from the past answers of our emminent
} } immunoglobulinologist Jan Leunissen that the size of an IgG is several
} } nm, I think more than 5nm.
} } Which means, in your case, that you should not expect more that a few
} } molecules per nanoparticle, except if they form multilayers.
} } As for the techniques, I wonder if a SEM could resolve this structure.
} } Although I have experience with analytical low res SEMs, I don't know
} } the limits of modern FEG SEMs.
} } I would be happy if some listers would care to share their opinion on
} } this matter.
} } It seems that AFM (atomic force microscopy) would be able to resolve
} } this structure, but I wonder if it would be possible in this condition
} } because AFM works on flat substrate and here you have round particles.
} } It would probably be possible to find some answers using cry-
} } microscopy, the same way scientists use it to resolve viral structures
} } at several anstrom resolution. It probably depends on how much time
} } and money your clients are wanting to spend and how critical this
} } information is for them.
} }
} } Regards,
} } Stephane
} }
} } ----- Original Message -----
} } X-from: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
} } To: nizets2-at-yahoo.com
} } Cc:
} } Sent: Friday, February 3, 2012 8:10 PM
} } Subject: [Microscopy] TEM of functionalized nanoparticles
} }
} }
} }
} }
} } ----------------------------------------------------------------------
} } ------ The Microscopy ListServer -- CoSponsor: The Microscopy Society
} } of America To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------
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} }
} } In recent years we have been getting requests to image functionalized
} } nanoparticles, such as 10 nm gold with proteins or peptides attached.
} } Generally the clients want to know how thick or large the organic
} } "shell" is, or how the protein/peptide is arranged on the surface.
} } Most of our experience has been with either inorganic particles (which
} } we view without staining) or larger organic structures such as viral
} } procapsids or large proteins (e.g., myosin) or protein assemblies,
} } which we negative stain. My questions for this list are as follows:
} }
} } - have others on the list had good luck imaging these kinds of
} } conjugates at the EM level, and if so, using what techniques?
} } - how much organic material must one have associated with the
} } nanoparticle to see it by negative staining or other methods? For
} } example, how hard is it to detect a 150 kD protein on a 10 nm gold
} } particle? An example would be IgG attached to a 10 nm gold particle.
} } - can you suggest any publications or review articles on the subject?
} } For some reason I'm not having much luck with my Pubmed search
} } strategies. A recent paper by Cao and Mao seems to deal mainly with
} } small gold particles on much larger protein aggregates, but perhaps
} } there are others out there.
} }
} } Many thanks (in advance) for your suggestions.
} }
} } Dr. Marie E. Cantino
} } Associate Professor of Physiology and Neurobiology Director, Electron
} } Microscopy Laboratory University of Connecticut, Unit 3242
} } Phone: 860-486-3588
} } Fax: 860-486-6369
} }
} }
} } ==============================Original
} } Headers==============================
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} } Stephane Nizet {nizets2-at-yahoo.com} 15, 42 -- Reply-To: Stephane Nizet
} } {nizets2-at-yahoo.com} 15, 42 -- Subject: Re: [Microscopy] TEM of
} } functionalized nanoparticles 15, 42 -- To: "marie.cantino-at-uconn.edu"
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}
} Dr. Marie E. Cantino
} Associate Professor of Physiology and Neurobiology Director, Electron
} Microscopy Laboratory University of Connecticut, Unit 3242 Phone:
} 860-486-3588
} Fax: 860-486-6369
}
}
} ==============================Original
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} -----Urspr�ngliche Nachricht-----
} Von: Mu� Wolfgang
} Gesendet: Montag, 06. Februar 2012 11:49
} An: 'mary.walker-at-csl.com.au'
} Cc: microscopy-at-microscopy.com
} Betreff: [Microscopy] Re: Counting virus particles

Dear Mary,
Greetings to Melbourne/Australia !

Hopefully you are not situated in the area of floodings!

Concerning your question(s) below I would like to point you to a MSA-
Listserver-thread (2007) with -/+ such background (a compilation of the
postings I could send to you relatively fast...but there would be also
other interesting questions and answers (which one can find also in
recent and a little bit "older" specific literature (from experts) on
that matter.
So, please, if you would like to get a compilation of the 2007-MSA-LS-
thread (instead of searching for them in the archives) please let me
know (and "allow" for sending to your mailbox...),

for now, best regards,
Wolfgang MUSS

} SALZBURG, AUSTRIA
}
} } -----Urspr�ngliche Nachricht-----
} } Von: microscopylistserver-noreply-at-microscopy.com
} } [mailto:microscopylistserver-noreply-at-microscopy.com]
} } Gesendet: Freitag, 03. Februar 2012 16:40
} } An: Mu� Wolfgang
} } Betreff: [Microscopy] Counting virus particles
}
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} } Email: mary.walker-at-csl.com.au
} } Name: mary walker
} } Organization: csl limited
} } Title-Subject: counting virus particles
} } Message:
} } Do any listers have experience with counting virus particles with
} latex
} } beads?
} } Is it possible to do better than order of magnitude?
} } What is the minimum number of particles per ml that can be counted?
} } What is the lower limit?
} }
} } thanks in advance
} }
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==============================Original Headers==============================
9, 37 -- From W.Muss-at-salk.at Mon Feb 6 11:34:15 2012
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Hi Marie,

Somewhere in the 90's I believe I read an abstract that showed excellent negative staining of gold particles attached to immunoglobulin. Unfortunately, I do not have access to that literature here so I can not help you with a reference. If I remember correctly in those studies first the Ig component was a adsorbed onto a grid film which was then, after rinsing incubated with the gold particles to study the way those particles bound and negatively stained. I believe these were proceedings from one of the international EM conferences, perhaps someone else has access?

Good luck,

Jan Leunissen

e: http://www.aurion.nl

On 4/02/2012, at 8:07 AM, marie.cantino-at-uconn.edu wrote:

}
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} In recent years we have been getting requests to image functionalized
} nanoparticles, such as 10 nm gold with proteins or peptides attached.
} Generally the clients want to know how thick or large the organic
} "shell" is, or how the protein/peptide is arranged on the surface.
} Most of our experience has been with either inorganic particles (which
} we view without staining) or larger organic structures such as viral
} procapsids or large proteins (e.g., myosin) or protein assemblies,
} which we negative stain. My questions for this list are as follows:
}
} - have others on the list had good luck imaging these kinds of
} conjugates at the EM level, and if so, using what techniques?
} - how much organic material must one have associated with the
} nanoparticle to see it by negative staining or other methods? For
} example, how hard is it to detect a 150 kD protein on a 10 nm gold
} particle? An example would be IgG attached to a 10 nm gold particle.
} - can you suggest any publications or review articles on the subject?
} For some reason I'm not having much luck with my Pubmed search
} strategies. A recent paper by Cao and Mao seems to deal mainly with
} small gold particles on much larger protein aggregates, but perhaps
} there are others out there.
}
} Many thanks (in advance) for your suggestions.
}
} Dr. Marie E. Cantino
} Associate Professor of Physiology and Neurobiology
} Director, Electron Microscopy Laboratory
} University of Connecticut, Unit 3242
} Phone: 860-486-3588
} Fax: 860-486-6369
}
}
} ==============================Original Headers==============================
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10, 21 -- From leunissen-at-aurion.nl Mon Feb 6 16:25:04 2012
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I was also amused by the "driving test" analogy and really smiling at
Steve Chapman's response.

I thought the old, Siemens Elmiskop had an great approach to an
operational mistake: a red bar would loudly clunk down into place, in
essence, screaming at the operator, "You made a BIG mistake, here!
Call someone who knows what they are doing." I know someone who nearly
fainted when this occurred (in the wee hours of the morning with no
one else in the building). I never had the chance to use the
instrument, though I really admired the high resolution images from
this excellent instrument.

--

John J. Bozzola, Ph.D.
Gleefully Retired Professor & Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

On Mon, Feb 6, 2012 at 4:56 PM, {protrain-at-emcourses.com} wrote:
} Hi All
}
} I am always amused when people talk about giving users a driving test before allowing them to use a microscope. Not only do I teach people to use electron microscopes I teach people to drive racing vehicles and it is only then that, with a smile, you realise just how much a driving test on a microscope differs dramatically from any other type of "driving test"! When you drive a car or any other vehicle and make an error you frighten yourself and perhaps worry that you will be hurt, and you do not make the same mistake again. Contrast this when there is no pain from making an error on a microscope. The best computing system that I have seen that trained people correctly was MS Dos and MS Word One, here make any error and you were faced with a screen stating "fatal error", the machine would then crash and all would be lost. You learned pretty quickly or just gave up!
}
} Yes people should undergo a microscope "driving test". The test should be designed to produce a result that clearly demonstrates that the operator took into account every pertinent feature of the instrument. From my experience a number of specimens are required each demanding an understanding of different features within the instrument. I am often forced to say, when evaluating laboratories and their staff, you may have 1nm instruments but do you realise you have 5nm staff? Staff testing in my mind is required and through repeated testing and under pressure the staff will develop their skills!
}
} Think of "Quality" the Japanese way - repeated evaluation and improvement.
}
} Some ramblings of an electron microscopist clocking up 48 years in the business.
}
} Steve
}
} Steve Chapman FRMS
} Senior Consultant Protrain
} For consultancy and training in electron microscopy world wide
} Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
} www.emcourses.com
}
} -----Original Message-----
} From: M. Redigolo [mailto:marcela.redigolo-at-mail.wvu.edu]
} Sent: 06 February 2012 17:53
} To: Allan Mitchell; Andrew D. Hollingsworth; Andrew Sullivan; Hands-Portman Ian; j.janssen-at-nki.nl; John Bozzola; Moore Jayma; Naomi McCallum; Patricia Nelson; protrain-at-emcourses.com; Richard E. Edelmann; sallystowe-at-gmail.com; Sherman Debra; Straszheim Warren E [BIOTC]; tommibarhorst-at-gmail.com; vray-at-partbeamsystech.com; Wim Hagen; Wim Van Den Broeck
} Subject: New management topic - discussion
}
} Dear all:
}
} During the benchmarking this past week, two topics came up that it seems
} many want to discuss.
} So, I'm putting the topics out there to know what is the common practice in
} other facilities.
}
} a) How to evaluate the efficiency (in terms of results, quality of data,
} budget, etc) of management, fees and users?
} b) Service contracts - self-service / all-inclusive / OEM service contract /
} labor-only OEM service contract / third-party
} service contract / per-hour paid service from OEM or third-party /
} self-service with OEM support or with third-party support?
}
} Any inputs on those topics?
}
} Currently here, we have full service contract (all-inclusive) for both JEOL
} instruments. In the case of the SEM, 5 years (3 years to go)
} and for the TEM, we pay it yearly. This is one point that I might review and
} change if I know that other model can also work fine and
} save us money.
}
} We do collect a copy of every publication resulted from using the facilities
} Users are very good in sending us these copies since we use
} them to justify increase of budget. Also, every user is requested to add an
} acknowledgement line mentioning the facilities, in every publication.
} Besides that, fees are reviewed yearly as I commented in the previous email.
} In terms of usage of the instruments, in some cases (as it is for the TEM)
} users undergo a "driver's test" before becoming unassisted users.
}
} OK. I promise I'll stop here and not crowd your inbox with long emails...
}
} Thanks again!
}
} Marcela.
}
}
}
} ------
} Dr. Marcela Redigolo
} Electron Microscopy Facility
} WVU Shared Research Facilities
} (304) 680-3007
} marcela.redigolo-at-mail.wvu.edu
}
}
}
}

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The Texas Society for Microscopy invites you to participate in
its 2012 meeting taking place on the TCU campus Apr. 12-14. The
meeting location is the Brown-Lupton University Union while the
host hotel will be the SpringHill Suites by Marriott University
Fort Worth. Shuttle service from the hotel to campus will be
available. The abstract deadline is March 1. Additional meeting
information and forms can be found on our website:
http://www.texasmicroscopy.org.

Workshops scheduled for Thursday, April 12th, 1 � 4 PM
Care & Feeding of Your Light Microscope and Light Source
Sample Preparation Techniques for Materials Science � Choosing
the Right Method by Robert Ranner, Product Manager, Leica
Applications Specialist
Etching, ion beam milling, slope cutting � what�s the difference?
Target surface preparation � getting it down to size.
Ultrasectioning � why go cryo?
Hands-on demonstration:
TXP � multi-functional target preparation device which can saw,
mill, grind and polish
TIC3X � automated ion beam milling system
UC7/FC7 � advanced room and cryo temperature ultramicrotome

Invited Speakers Friday, April 13, 2012
Dr. Ray H. Baughman, Alan G. MacDiarmid NanoTech Institute, The
University of Texas at Dallas
- Biscrolling Nanofiber Sheets and Functional Guests into
Multifunctional Yarns for Energy Applications

Dr. Aydogan Ozcan, Bio- and Nano-Photonics Laboratory, UCLA
� Photonics-based Telemedicine Technologies Toward Smart Global
Health Systems

==============================Original Headers==============================
4, 27 -- From r-holdford-at-ti.com Mon Feb 6 19:05:39 2012
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Ah, fond memories. My first real experience in TEM (besides playing with a
Philips 75 as an undergrad) was in the lab of Humberto Fernandez-Moran at
the University of Chicago. We had Siemens Elmiskop 1 and 1a microscopes.
Oh yes...saw that red bar on numerous occasions. The scope had a mechanical
valve block and if it was not turned properly the bar sprang up. Made enough
noise so those in the next room knew that you did a no-no.

However, I still have glass negatives of some images taken from that
instrument. I have never seen better coming from any modern W-filament
instrument. Check out the T4 phage (at www.dsimagingllc.com under "Images }
animal"), which was taken 45 years ago when I had been using the Siemens
Elmiskop I for about 6 months. Before being able to look at real sample, we
had to produce a perfect objective stigmation series on a hole sample at
very high mag, using the mechanical stigmator system available in those
days. I really learned how to stigmate and focus incredibly well with almost
no illumination. It has been a skill that I still value extremely highly.
Spent a lot of time in the dark working on that microscope along with a
young grad student who became my husband. There is a lot to be said for
spending time in the dark! One big advantage we had was using what I
believe were the first pointed filaments that were made in the lab. But that
is another story. For those calculating my age, I was very young at the time
to have access to such an instrument.....

Debby Sherman

On 2/6/12 6:16 PM, "John Bozzola, Phd. Director" {bozzola-at-siu.edu} wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} I was also amused by the "driving test" analogy and really smiling at
} Steve Chapman's response.
}
} I thought the old, Siemens Elmiskop had an great approach to an
} operational mistake: a red bar would loudly clunk down into place, in
} essence, screaming at the operator, "You made a BIG mistake, here!
} Call someone who knows what they are doing." I know someone who nearly
} fainted when this occurred (in the wee hours of the morning with no
} one else in the building). I never had the chance to use the
} instrument, though I really admired the high resolution images from
} this excellent instrument.

==============================Original Headers==============================
9, 31 -- From dsherman-at-purdue.edu Mon Feb 6 19:46:52 2012
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I imagine you would have difficulty using AFM if you needed to count things attached to particles. AFM gives you a 2D projection of the sample, you would miss anything under or "shadowed" by the particle or anything else. AFM doesn't do reentrant profiles (typically).
If all you need is to see if something's attached, get some size information (caution about reentrant profiles again), and you have well characterized particles to attach to, AFM might be able to help you.

Jonathan Abbott

On Feb 6, 2012, at 7:48, marie.cantino-at-uconn.edu wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi Stefan,
}
} I do understand the difference between mass and length.
}
} The problem is that the clients usually can only give us the molecular
} weight and (if we're lucky) the number of molecules per particle. If
} they knew how the proteins/peptides were arranged or the linear
} dimensions of the coating they wouldn't need to come to me. What I'm
} trying to find out from the listserver or the literature is whether we
} can reasonably expect to be able to detect a medium sized protein such
} as IgG on an electron dense gold particle, given the various artifacts
} of techniques such as negative staining.
}
} We only have TEM (w/a tungsten emitter) and FESEM in our facility, but
} I can send them elsewhere for AFM.
}
} Marie
}
} On Feb 6, 2012, at 5:33 AM, nizets2-at-yahoo.com wrote:
}
} }
} }
} }
} } ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } America
} } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------------
} }
} } Hi Marie,
} }
} } I like these projects which put the techniques to their limits.
} } I think the reason why you have difficulties finding publications on
} } this matter and that the question is not trivial at all.
} } First of all, and to start with a relaxed note, the mixing of kDa
} } und nm in one sentence made me smile because it makes as much sense
} } as asking if 1kg of something fits in one meter.
} } The Dalton is a unit of mass and does not give a good indication of
} } the dimension or the shape of a molecule.
} } That said, I seem to remember from the past answers of our emminent
} } immunoglobulinologist Jan Leunissen that the size of an IgG is
} } several nm, I think more than 5nm.
} } Which means, in your case, that you should not expect more that a
} } few molecules per nanoparticle, except if they form multilayers.
} } As for the techniques, I wonder if a SEM could resolve this
} } structure. Although I have experience with analytical low res SEMs,
} } I don't know the limits of modern FEG SEMs.
} } I would be happy if some listers would care to share their opinion
} } on this matter.
} } It seems that AFM (atomic force microscopy) would be able to resolve
} } this structure, but I wonder if it would be possible in this
} } condition because AFM works on flat substrate and here you have
} } round particles.
} } It would probably be possible to find some answers using cry-
} } microscopy, the same way scientists use it to resolve viral
} } structures at several anstrom resolution. It probably depends on how
} } much time and money your clients are wanting to spend and how
} } critical this information is for them.
} }
} } Regards,
} } Stephane
} }
} } ----- Original Message -----
} } X-from: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
} } To: nizets2-at-yahoo.com
} } Cc:
} } Sent: Friday, February 3, 2012 8:10 PM
} } Subject: [Microscopy] TEM of functionalized nanoparticles
} }
} }
} }
} }
} } ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } America
} } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------------
} }
} } In recent years we have been getting requests to image functionalized
} } nanoparticles, such as 10 nm gold with proteins or peptides attached.
} } Generally the clients want to know how thick or large the organic
} } "shell" is, or how the protein/peptide is arranged on the surface.
} } Most of our experience has been with either inorganic particles (which
} } we view without staining) or larger organic structures such as viral
} } procapsids or large proteins (e.g., myosin) or protein assemblies,
} } which we negative stain. My questions for this list are as follows:
} }
} } - have others on the list had good luck imaging these kinds of
} } conjugates at the EM level, and if so, using what techniques?
} } - how much organic material must one have associated with the
} } nanoparticle to see it by negative staining or other methods? For
} } example, how hard is it to detect a 150 kD protein on a 10 nm gold
} } particle? An example would be IgG attached to a 10 nm gold particle.
} } - can you suggest any publications or review articles on the subject?
} } For some reason I'm not having much luck with my Pubmed search
} } strategies. A recent paper by Cao and Mao seems to deal mainly with
} } small gold particles on much larger protein aggregates, but perhaps
} } there are others out there.
} }
} } Many thanks (in advance) for your suggestions.
} }
} } Dr. Marie E. Cantino
} } Associate Professor of Physiology and Neurobiology
} } Director, Electron Microscopy Laboratory
} } University of Connecticut, Unit 3242
} } Phone: 860-486-3588
} } Fax: 860-486-6369
} }
} }
} } ==============================Original
} } Headers==============================
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} } 5, 18 -- Received: from mta1.uits.uconn.edu (mailscanner.uconn.edu
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} } 5, 18 -- From: Marie Cantino {marie.cantino-at-uconn.edu}
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} } 5, 18 -- Subject: TEM of functionalized nanoparticles
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} } nanoparticles
} } 15, 42 -- To: "marie.cantino-at-uconn.edu" {marie.cantino-at-uconn.edu}
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}
} Dr. Marie E. Cantino
} Associate Professor of Physiology and Neurobiology
} Director, Electron Microscopy Laboratory
} University of Connecticut, Unit 3242
} Phone: 860-486-3588
} Fax: 860-486-6369
}
}
} ==============================Original Headers==============================
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I had a session on a Philips CM20 today.� I was working at relatively high
mag and I wanted to shift the image instead of using the mechanical stage
shifts.� For the life of me, I could not figure out how to do it.� I�m not
that familiar with the instrument, but I�m sure that it can be done.� Can
anyone tell me the controls for it?

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

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9, 36 -- Subject: image shift on CM20
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let me briefly comment on one sentence in Jonathan's comment from yesterday.

--
Prof. Dr. Reinhard Rachel
University of Regensburg
Centre for EM / Anatomy
Faculty of Biology & Preclin. Med.
Universitaetsstrasse 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-regensburg.de
office: VKL 3.1.29

next microscopy conferences:
http://www.emc2012.org.uk/
EMC 2012 in Manchester, UK
http://www.mc2013.de/
MC2013 in Regensburg, Germany

} } }
} I imagine you would have difficulty using AFM if you needed to count things
} attached to particles.
} AFM gives you a 2D projection of the sample,

no, this is not the case. AFM will give you an image representing the surface / surface relief of the sample, but not a 2D projection (in the sense as the TEM does). You will not get any information of the internal parts of the sample.

} you would miss anything under or "shadowed" by the particle or anything else. AFM
} doesn't do reentrant profiles (typically).

} If all you need is to see if something's attached, get some size information
} (caution about reentrant profiles again), and you have well characterized
} particles to attach to, AFM might be able to help you.

here, I fully agree.
Kind regards,
Reinhard

==============================Original Headers==============================
11, 25 -- From reinhard.rachel-at-biologie.uni-regensburg.de Tue Feb 7 03:10:45 2012
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Hello Amit,

"stripes" in images are often the result of interference. It often helps to
determine the frequency to eliminate sources (for example if the frequency
is 50 or 60 Hz, it usually comes from a power source). Have you tried to
determine the frequency?

Mike Bode, Ph.D.
ResAlta Research Technologies Corp.

2102 Beech Ct
Golden, CO 80401
USA

p: +1-303-234-9270
f: +1-303-202-6350
Mike.Bode-at-ResAltaTech.com
www.ResAltaTech.com

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Email: amit.welcomes.u-at-gmail.com Name: Amit

Organization: Indian institute of science

Title-Subject: [Filtered] Diagonal bands in STEM mode

Message: we recently installed JEOL JEM-2100F microscope. rest is
working fine but in STEM mode there are Diagonal Bands of alternate
brightness coming(all bands or strips are of same thickness, brightness
and contrast). on increasing scan time band's thickness first increases
then at even higher scan times it decreases. Engineers are on it but
cant pin point the fault. i was wondering if anyone encountered it
before or knows why this is happening

Login Host: 14.139.128.11
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Regarding�AFM of gold nanoparticles with proteins attached, I have some experience with�this.

As was�mentioned, AFM (high resolution at least) works best with flat samples. You can
certainly images round particles with no difficulty, but you will not normally�get many details. What you will see is a �blob�, most likely. Now, if your�particles are nice (i.e. close to monodisperse), and your protein size is�significant compared to your gold particle size, what you can do, is compare the��unfunctionalised� size (height of the blob) to the functionalised size, and�use this difference as a measure of how much, if any, protein got attached. AFM�has very good resolution in height, getting images good enough to make these�measurements would be simple. The limiting factor here will most likely be,�what is the significance of your height measurements, given the range of height�of the �bare� particles. Also, how many AFM images you can bear to measure, to�get� a significant sample.

�

By the way,�no, we don�t directly compare kDa and nanometres, but I understood exactly why�the poster said this...ask a biochemist how big their protein is, and they will always reply in terms of Daltons. We can, however make a *guess* at the size based on the�height, and I can tell you that your 150kDa particle WILL make a significant difference�to the 10 nm AuNP�s diameter, since I have done this with much smaller proteins.

�

On the�other hand, there is no doubt that if the negative staining works well, and you�can image both the the AuNPs and proteins, it will be a nicer result, and probably less�time consuming.

�

Good Luck!

Pete.

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Thank you, Wim. I believe this is what I want. I didn't use the HR-TEM
mode, I used the TEM mode. Wim Hagen's direct email message to me included
a readout image of the HR-TEM page and it shows the Image Shift as well as
some other HR-TEM functions. His answer was to use the HR-TEM mode, choose
Parameter page and the appropriate control pages come up.

Chad Parish, John Minter, and Reinhard Rachel suggested using the
Alignment-Image Shift. I did try that and all it did when I touched the
multi-knobs was to beep at me. John said that this function is for keeping
the different mags aligned.

Thank you all for your help. Next time I'm on the microscope, I'll try it.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: Wim Hagen [mailto:wim.hagen-at-me.com]
Sent: Tuesday, February 07, 2012 12:28 AM
To: S.Walck-at-cox.net

Another thing you could try is high-angle Pt/C shadowing.

I don't know if it's easier to find an AFM or a working high vacuum evaporator these days, though!

Andy Bowling

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I already gave my opinion to Marie about this offline but since there were several answers in the list going in this direction I would like to share some thoughts with the list.

Negative staining may work. This means: you may see your proteins, you may get a result. You may even get a nice picture.
But hey, are we artists or are we scientists?
Our job is not to get nice pictures, it is to get meaningful pictures.

Marie wants to know how the antibodies (or other proteins) are organized around gold particles in a matrix, most probably water or aqueous solution.
I am not sure the best way to know that is by drying the sample with a heavy stain.
Do you really think it may not interfere with the binding forces between the antibody and the gold particle? I wouldn't bet a cent on it!

Regards,
Stephane

----- Original Message -----
X-from: "petereaton-at-hotmail.com" {petereaton-at-hotmail.com}
To: nizets2-at-yahoo.com
Cc:
Sent: Tuesday, February 7, 2012 6:33 PM

�

On the�other hand, there is no doubt that if the negative staining works well, and you�can image both the the AuNPs and proteins, it will be a nicer result, and probably less�time consuming.

�

Good Luck!

Pete.
��

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Hello Stephane,

This conundrum you indicate is like life. One picks his or her poison according to the needs, goals, facilities available, time, and economic/collaborative capabilities. Sometimes an association is all the information that is needed to answer a question. All of the methods available today for visualizing proteins have flaws. Negative stain EM is a good first start with other techniques such as cryo-EM and AFM later based on what was learned by "crude" negative stain EM, if indeed one is attempting to study how proteins are organized around a matrix.

Best regards,

Charles Humphrey, Ph.D.
CDC
Atlanta, GA 30333

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Wednesday, February 08, 2012 5:28 AM
To: Humphrey, Charles (CDC/OID/NCEZID)

I already gave my opinion to Marie about this offline but since there were several answers in the list going in this direction I would like to share some thoughts with the list.

Negative staining may work. This means: you may see your proteins, you may get a result. You may even get a nice picture.
But hey, are we artists or are we scientists?
Our job is not to get nice pictures, it is to get meaningful pictures.

Marie wants to know how the antibodies (or other proteins) are organized around gold particles in a matrix, most probably water or aqueous solution.
I am not sure the best way to know that is by drying the sample with a heavy stain.
Do you really think it may not interfere with the binding forces between the antibody and the gold particle? I wouldn't bet a cent on it!

Regards,
Stephane

----- Original Message -----
X-from: "petereaton-at-hotmail.com" {petereaton-at-hotmail.com}
To: nizets2-at-yahoo.com
Cc:
Sent: Tuesday, February 7, 2012 6:33 PM

�

On the�other hand, there is no doubt that if the negative staining works well, and you�can image both the the AuNPs and proteins, it will be a nicer result, and probably less�time consuming.

�

Good Luck!

Pete.
��

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Hello list!
�
We have taken almost all the shielding from our Tecnai G20 down in order to make some repairs and now that I am ready to mount them again, I ask myself:
Should I ask FEI to come and control the X-radiation?
We work mostly at 100-120kV but sometimes we use 200kV so X-rays should be considered, but is there a risk of leak after unmounting/remounting the shielding?
Thanks in advance for sharing your thoughts.

Regards,
Stephane

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Thanks to everyone who has responded to my question. Since there
seemed to be quite a bit of interest, I will summarize what I've
discovered so far (I hope those who responded off-line will not mind
being paraphrased, and my apologies if I accidentally omitted your
suggestions. There were a lot of emails.)

Several people suggested publications, and I did additional searches
on my own (though not exhaustive). Google Scholar proved to be more
useful than Pubmed, perhaps because many of these papers are in
chemistry journals.

Here are some papers that include EM methods (noted in parentheses),
in no particular order:

Transmission Electron Microscopy as a Tool to Image Bioinorganic
Nanohybrids: The Case of Phage-Gold Nanocomposites MICROSCOPY RESEARCH
AND TECHNIQUE 74:627�635 (2011) (negative staining, mainly of larger
protein assemblies)

Apoferritin-Templated Synthesis of Encoded Metallic Phosphate
Nanoparticle Tags. Anal. Chem. 2007, 79, 5614-5619. (TEM unstained and
negative stained).

Direct Imaging of Soft-Hard Interfaces Enabled by Graphene. Nano
Letters 2009 Vol 9 No 9 3365-3369 (very cool images using high
resolution TEM and substrate, unstained)

In Vivo Imaging of Quantum Dots Encapsulated in Phospholipid
Micelles. SCIENCE VOL 298 29 NOVEMBER 2002 1759-1762 (negative
stained)

Electron Density Imaging of Protein Films on Gold-Particle Surfaces
With Transmission Electron Microscopy. Ludger Ju�rgens,* Alfons
Nichtl, and Ulf Werner. Cytometry 37:87�92 (1999) (TEM, unstained and
ruthenium stained prior to conjugation)

Controlled Encapsidation of Gold Nanoparticles by a Viral Protein
Shell. LiNa Loo, Richard H. Guenther, Veronica R. Basnayake, Steven
A. Lommel, and Stefan Franzen*. J. AM. CHEM. SOC. 2006, 128,
4502-4503 (negative stain TEM and possibly cryo, but results not shown)

A fluorescent, paramagnetic and PEGylated gold/silica nanoparticle for
MRI, CT and fluorescence imaging Matti M. van Schoonevelda, David P.
Cormodeb, Rolf Koole, J. Timon van Wijngaardena, Claudia Calcagno,
Torjus Skajaa, Jan Hilhorst, Dannis C. �t Hartc, Zahi A. Fayadb,
Willem J. M. Mulder* and Andries Meijerinka. Contrast Media Mol.
Imaging 2010, 5 231�236 2010. (negative and positive stain)

DNA-Origami-Directed Self-Assembly of Discrete Silver-Nanoparticle
Architectures. Suchetan Pal, Zhengtao Deng, Baoquan Ding, Hao Yan,*
and Yan Liu. Angew. Chem. 2010, 122, 2760 �2764 (negative stain w/
uranyl formate and unstained with STEM; not typical gold
nanoparticles, but pretty interesting structures!)

New Frontiers in Gold Labeling. James F. Hainfeld and Richard D.
Powell The Journal of Histochemistry & Cytochemistry 48(4): 471�480,
2000 (STEM images of nanogold labelled Fab)

Electron microscopy localization and characterization of
functionalized composite organic-inorganic SERS nanoparticles on
leukemia cells Ai Leen Koh , Catherine M. Shachaf, Sailaja Elchuri,
Garry P. Nolan, Robert Sinclair. Ultramicroscopy 109 (2008) 111�121
(negative stain TEM, SEM, BSE, Quantomix SEM and EDS)

In Vivo Imaging of Quantum Dots Encapsulated in Phospholipid Micelles
Benoit Dubertret, Paris Skourides, David J. Norris, Vincent Noireaux,
Ali H. Brivanlou, Albert Libchaber. SCIENCE VOL 298 29 NOVEMBER2002
(negative stain TEM)

Methods suggested:

-Negative staining, Uranyl acetate (0.5-2%) or ammonium molybdate
(1-2% w/trahalose), preferably with FEG or LaB6. Use
unfunctionalized particles as a control. Use of trehalose or an
embedding medium such as methyl cellulose was also suggested (see
Methods in Molecular Biology 369 Electron Microscopy: Mehods and
Protocols, 2nd edition, Chapter 7 by Robin Harris). In general,
drying artifacts such as flattening are likely to be observed with
negative staining, but use of these embedments will reduce this.

-Use of graphene support films see paper above in Nano Letters.

-dynamic light scattering

-cryo TEM or cryo FESEM were suggested

-Pt/C shadowing was suggested

-AFM. The dimension of the coating can be estimated by comparing the
height of the coated and uncoated particles.

Dr. Marie E. Cantino
Associate Professor of Physiology and Neurobiology
Director, Electron Microscopy Laboratory
University of Connecticut, Unit 3242
Phone: 860-486-3588
Fax: 860-486-6369

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Hello,

Our client wants us to image proteins using SEM. The protein is dissolved in water and currently stored at -20degrees C. What do you suggest? This is what I'm thinking:

Float a formvar-coated grid on drop of solution or place a drop of solution on a poly-L-lysine cover slip.

Fix grid or cover slip.

Rinse.

Osmicate grid.

Rinse and dry.

Sputter coat.

Thank you very much,

Jaci

Jaclynn Lett
Senior Research Technician
Research Center for Auditory and Visual Studies
Washington University School Of Medicine
Saint Louis, Missouri
lettj-at-ent.wustl.edu

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In my quest for an affordable printer to make B+W images of carbon black particles, I floated this question to the microscopy list server: "What's going to give me the nicest B+W TEM print?" After all, the images included in a report may be all the client knows about you.

I got a few interesting answers and in the process learned a bit about printers. But first, here's what I'm currently using: an HP2550 with copier paper. By the way the noise as stopped and the printer continues to run fine.

Here's what was suggested to me:
HP C5180,
HP2300,
HP8750,
Epson 800,
Sony Medical Printers.

Most people felt a color printer gave better gray scale images than a simple B+W printer but I was cautioned that a single black ink cartridge would be unable to create the entire tonal shading that good B+W prints need. Two different black ink cartridges could accomplish this.

Everyone had their own favorite printer and of course visiting vendor websites at 72 lines per inch of monitor resolution makes everyone's images look good. The best advice I got was to print images on as many printers as possible and then decide.

As a personal note, I'm not going to shop for printers right now, but I will try a ream of a better paper. Since several people print both text and images on our printer, running photo quality paper isn't an answer we can afford.

Increasing the resolution of our TEM camera would also be a big help,......

Thanks again for everyone's advice.

Frank

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HI Philip-
I have tried knives from different sources but have always come back to
Diatome. New or resharpened, they are great. The HIsto knife is
wonderful for 0.5-2 um sections. I haven't used glass in years. The
ultras are great from end-to-end. You also get great customer support
from both the US and Swiss offices.
Lee Cohen-Gould, M.S., C.E.M.T.
Chair, MSA Certification Board

Sr. Staff Associate in Biochemistry and
Cell & Developmental Biology
Director, Electron Microscopy & Histology and Optical Microscopy Core
Facilities Weill Cornell Medical College

lcgould-at-med.cornell.edu
voice (212)746-6146
fax (212)746-8175
http://www.med.cornell.edu/research/rea_sup/
http://www.cornellbiochem.org
http://www.cornellcelldevbiology.org

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Email: slakmon-at-yahoo.com Name: Slakmon

Title-Subject: [Filtered] Diamond knives & cleaning methods

Message: Good Afternoon,

I was hoping to get people's feedback and opinion on the different
diamond knives on the market. Your opinion on new, resharpened would be
helpful as well. What methods that you found best suited in cleaning
your knives without damaging your knives. Please feel free to respond on
the group
or offline directly to my email address.

Thank you in advance,

Philip

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http://www.biotech.ufl.edu/EM/data/viruscount.html

Try the above link for help.

Cheers,

FC Monson

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Email: mary.walker-at-csl.com.au Name: mary walker

Organization: csl limited

Title-Subject: [Filtered] counting virus particles

Message: Do any listers have experience with counting virus particles
with latex beads?
Is it possible to do better than order of magnitude?
What is the minimum number of particles per ml that can be counted?
What is the lower limit?

thanks in advance

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Hello Mary,

I have done my fair share of virus particle counts over too many decades
and have some opinions about the process. Latex beads may be used in
counting but I would not recommend them as a standard except as a rough
size standard. The manufacturer of the beads would agree as well I
think. It is a supreme leap of faith to believe that beads adsorb and
remain adsorbed to formvar-carbon films or any other films in the same
manner as viruses. I also expect different viruses passively adsorb
differently as well. If you are using airfuge techniques the adsorption
issue may be somewhat improved but one still has to stain and assume
beads remain adsorbed to the grids exactly as the viruses. I know there
are many publications that have used latex beads as counting standards
but my understanding of viruses causes me to be skeptical of including
latex beads for counting. One can count the viruses directly using the
grid squares as a constant and then extrapolate, or even better with
digital images a digital image grid of precise size can be overlaid and
is helpful.

It is also risky to consider reliability beyond 10 -fold. My experience
is that there is considerable variability within the 10-fold limits. I
suspect this is true with most counting methods; EM or otherwise. The
major advantage with EM is that one can actually see aggregates when
they occur (another issue with virus counts if other counting methods
are used).

If one assumes on a 300 mesh grid with a grid hole of 61 microns, 20
particles within that grid square would represent approximately 107
particles per ml. Therefore, considering the fields one would have to
view and identify particles, then probably 106 particles per ml is about
the limit one would wish to search for counting. This is what I would
consider as a lower practical limit. One could improve on the lower
limit by looking at many-many grid squares but that would require
considerable searching, patience, and time.

I hope this is helpful; if you have additional questions I may be
reached by email or my address below.

Happy counting,

Charles Humphrey
Mailstop G32
US Centers for Disease Control and Prevention (CDC)
Atlanta, GA 30333

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Title-Subject: [Filtered] counting virus particles

Message: Do any listers have experience with counting virus particles
with latex beads?
Is it possible to do better than order of magnitude?
What is the minimum number of particles per ml that can be counted?
What is the lower limit?

thanks in advance

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Hi Ravi,

It could be that your sample was not at the 'standard focus' height. Set
your objective lens to the 'standard focus' value and then move your
sample height to get to focus.

I hope this helps.

Ke-Bin

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} Email: ravi.thakkar369-at-gmail.com Name: Ravindra Thakkar
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} Title-Subject: [Filtered] TEM : Focusing Problem.
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Dear All,
I am looking for someone old enough to be fit in Visual Basic 6 ;-)
to help me with an existing VB6 program to remote my Philips 525 SEM.
Knowledge of German language would be perfect...
If interested, please contact me offline.

Best wishes,
Stefan
--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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My experience with the Histo knife is similar. Its an incredible knife!

Dr. Gary B. Carr
Pacific Endodontic Research Foundation
San Diego, CA 92121
----- Original Message ----- } From:
{microscopylistserver-noreply-at-microscopy.com}
To: {gary-at-perfendo.com}
Sent: Wednesday, February 08, 2012 3:08 PM

I would request that you respond on-line. I would be interested in others'
views.

Peggy Sherwood
Lab Associate, Photopathology
Wellman Center for Photomedicine (EDR 214)
Massachusetts General Hospital
50 Blossom Street
Boston, MA 02114-2696
617-724-4839 (voice mail)
617-726-6983 (lab)
617-726-1206 (fax)
msherwood-at-partners.org

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Email: slakmon-at-yahoo.com Name: Slakmon

Title-Subject: [Filtered] Diamond knives & cleaning methods

Message: Good Afternoon,

I was hoping to get people's feedback and opinion on the different
diamond knives on the market. Your opinion on new, resharpened would be
helpful as well. What methods that you found best suited in cleaning
your knives without damaging your knives. Please feel free to respond on
the group
or offline directly to my email address.

Thank you in advance,

Philip

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I'd have thought the osmium would be an unnecessary step since you're
coating the specimen before viewing. To fix the protein I'd be inclined
to use glutaraldehyde vapour - we've fixed problematic complexes before
by blotting the grid then holding it in the air space in a bottle of the
fixative to ten to thirty seconds. After a quick rinse you ought to be
able to dry it and put straight in the coater.
Cheers
Ian

http://www2.warwick.ac.uk/fac/sci/lifesci/facilities/imaging

-----Original Message-----
X-from: LettJ-at-ent.wustl.edu [mailto:LettJ-at-ent.wustl.edu]
Sent: 08 February 2012 19:54
To: Hands-Portman, Ian

Hello,

Our client wants us to image proteins using SEM. The protein is
dissolved in water and currently stored at -20degrees C. What do you
suggest? This is what I'm thinking:

Float a formvar-coated grid on drop of solution or place a drop of
solution on a poly-L-lysine cover slip.

Fix grid or cover slip.

Rinse.

Osmicate grid.

Rinse and dry.

Sputter coat.

Thank you very much,

Jaci

Jaclynn Lett
Senior Research Technician
Research Center for Auditory and Visual Studies Washington University
School Of Medicine Saint Louis, Missouri lettj-at-ent.wustl.edu

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Email: tomw-at-uidaho.edu Name: Tom Williams

Organization: University of Idaho

Title-Subject: [Filtered] Carbon Coater for TEM and FESEM
Message: Greetings.

I have decided that it is time to replace our "Classic" Varian
evaporative coater with a newer model. A day of replacing vacuum hoses,
and finding an inventory tag labeled: "Property of the Manhattan
Project," pushed me over the absorption edge.

Request: recommendations for a replacement unit? We operate a FESEM
and a 200kV TEM housed in a campus-wide Core Facility with a varied
client list (minerals, polymers, inorganics, and biologicals). We need
a coater suitable for routine SEM samples, EDS, hi-res FESEM work, TEM
grids, etc. so I'm looking for a relatively high-performance turbo
pumped system. I am prepared to expend a reasonable amount of capital
for this.

I would appreciate recommendations and/or suggestions for models,
vendors, etc.
Although I welcome contacts by vendors directly, I am looking for input
from end-users. Ranting diatribes are welcome as long as they are
entertaining!

Thanks,
Tom Williams

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I would suggest that it would be very important to get either the manufacturer's engineers or someone who is trained to that standard to do the re-assembly and x-ray check afterwards using the appropriate type of meter rather than any old geiger counter. Most of the training I had in the early days emphasised the need for scrupulous checks after a major re-assembly or modification and it was even more important for 100Kv+.

In the olden days we had a Siemens IA that spectaculalrly failed radiation checks when operated at 100kv but was fine at the lower voltages.

Hope this helps.

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: nizets2-at-yahoo.com [nizets2-at-yahoo.com]
Sent: 08 February 2012 16:07
To: Malcolm Haswell

Hello list!

We have taken almost all the shielding from our Tecnai G20 down in order to make some repairs and now that I am ready to mount them again, I ask myself:
Should I ask FEI to come and control the X-radiation?
We work mostly at 100-120kV but sometimes we use 200kV so X-rays should be considered, but is there a risk of leak after unmounting/remounting the shielding?
Thanks in advance for sharing your thoughts.

Regards,
Stephane

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12, 37 -- "Microscopy MSA
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12, 37 -- Subject: RE: [Microscopy] X ray control after dismounting a TEM
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Many thanks for taking the time to share your thoughts with me, I appreciated it very much!

The majority (5 versus 2) thinks that, although the risk is minimal, it would be well advised to control the X-ray leakage of the system after remounting.
One person mentioned the opening the column which I think is a good point (actually she talked about breaking the column but I don't think I am strong enough).
I didn't say it but we didn't open the column, in which case I think it is very important to check the X-ray leakage after remounting.

Have a nice and safe day

Stephane

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Dear Philip, dear all,

since Peggy requested "on-line responses I will do this.

Concerning
Q#1a: Diamond knife type: I am seconding (just as a really satisfied USER)
the opinions of Dr. Lee Cohen-Gould, Dr. Gary B. Carr as posted before.
I too have tested in my long life as EM-ist several types (but it seems that
today there are only few other companies dealing with Diamond knives
(ultra- as well as so called Histo-Type).
I know several 10 -20 years ago there were trials to push the Sapphire knife types
as a good alternative to the costly/expensive diamond knives, but have never found
real reports on their "life" / use (neither personal notes or in publications).

To be fair, since we previously have read about DIATOME knives, I honestly would
like to add the following here:
I am aware of the recent efforts of
{ Scimed GmbH
Scientific and Medical Products
Am Muehlenkanal 23
D-46419 Isselburg
Germany
Office: +49 2874 9055833
Mobile: +49 170 5121862
Fax: +49 2874 901495
Private: +49 2874 901494
info-at-dEYEmond.eu
www.dEYEmond.eu
skype: scimed_detlef }

to get into the market. (Disclaimer: No affiliation with, NO FINANCIAL INTEREST)

Unfortunately I was not able to test their new brands of Deyemond-knives
(ultra-, as well as Histo-Type, and forthcoming: cryo-knives) in comparison
with the Diatome knives I use.

Q#1b: Cost-efficiency: comparing costs of a good diamond knife with the costs
of the raw material(s) for fabricating knives (1) Glass stripes, 2) knife breaker
3) sticking tapes, 4) dental wax or plastic trough-boats and 5) TIME to make the
glass knives, 6) material and time lost in fabricating low quality cutting edges)
it seems to me that you have invested well in/with a diamond knife.
If treated properly the yield of a new diamond knife is 3000-5000 sections
(or more, the Histo-type for big section areas = 3/5 - 5/6 mm perhaps
depending on the (harder) contents in the specimen something less).
I also have (fortunately) never used glass knives since 1985 and usually
resharpening happens (needed) once in two years (unless there has been an
"accident" to the cutting edge).

Q#2: Cleaning: every knife comes in its original delivery box, containing
also polystyrol (foamy-) rods instead of former used rods from elder (?).
Also enclosed (and to be found on the web or via the company service)
instructions how to clean (and keep clean) your diamond knife.
I don't think that you can make mistakes if you follow strictly those
instructions quite straightly/strictly and if you would like to instruct
your staff or yourself you will get also a wall poster on that
(cleaning ONLY with A. dest. or Ethanol (on the polystyrol rod), if there
were heavy resistant remnants of plastic you incubate the knive over night
in A.dest. and try again with the rod.)

Personally I greatly should acknowledge some informations on the types
(brands, eventually dealing /offering company) of
diamond knives used these times out there...
so Google can be off this time...(;-))

Just my 2 cents,

Wolfgang MUSS
SALZBURG, Austria

} -----Urspr�ngliche Nachricht-----
} Von: microscopylistserver-noreply-at-microscopy.com
} [mailto:microscopylistserver-noreply-at-microscopy.com]
} Gesendet: Donnerstag, 09. Februar 2012 09:24
} An: Mu� Wolfgang
} Betreff: [Microscopy] Re: Diamond knives [also resharpened] & cleaning
} methods
}
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} I would request that you respond on-line. I would be interested in
} others'
} views.
}
}
}
} Peggy Sherwood
} Lab Associate, Photopathology
} Wellman Center for Photomedicine (EDR 214)
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} [mailto:microscopylistserver-noreply-at-microscopy.com] Sent: Wednesday,
} February 08, 2012 4:45 PM
} To: Sherwood, Margaret
} Subject: [Microscopy] Diamond knives cleaning methods
}
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} Title-Subject: [Filtered] Diamond knives & cleaning methods
}
} Message: Good Afternoon,
}
} I was hoping to get people's feedback and opinion on the different
} diamond knives on the market. Your opinion on new, resharpened would be
} helpful as well. What methods that you found best suited in cleaning
} your knives without damaging your knives. Please feel free to respond
} on
} the group
} or offline directly to my email address.
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} Thank you in advance,
}
} Philip
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Also put out on the confocal listserver

Dear Colleagues,

I have to take photographs of GFP newborn and adult mice to support a microscopy paper. This is to show GFP throughout the coat of the newborns, and in the tail tips and feet of the adults.

Normally mice are photographed with a 60mm f# 2.8 lens on a Nikon D70S with sync flash at 1/60th or 1/125th sec with an ISO of 100 or 200 set on the camera CCD. That, however, is for normal room lighting – plenty of photons.

We don’t want to go above ISO 800 because of noise, yet need to freeze the movement of the mice.

Has anyone done this, and can advise on exposure, whether to use sync flash as well as the UV source.

I have been advised to anaesthetise the mice and photograph out of the cage. We'd prefer not to anaesthetise, but there may be no option.

Any advice please, or if you can direct me to someone who has done this. All input gratefully received.

Thanks,
Jeremy Sanderson
Bio-Imaging, MRC Harwell

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I second that. I have used a histo knife for years. Wouldn't go back
to glass
ever!

Peggy Sherwood
Lab Associate, Photopathology
Wellman Center for Photomedicine (EDR 214)
Massachusetts General Hospital
50 Blossom Street
Boston, MA 02114-2696
617-724-4839 (voice mail)
617-726-6983 (lab)
617-726-1206 (fax)
msherwood-at-partners.org

-----Original Message-----
} From: microscopylistserver-noreply-at-microscopy.com
[mailto:microscopylistserver-noreply-at-microscopy.com] Sent: Wednesday,
February 08, 2012 6:12 PM
To: Sherwood, Margaret

HI Philip-
I have tried knives from different sources but have always come back to
Diatome. New or resharpened, they are great. The HIsto knife is
wonderful for 0.5-2 um sections. I haven't used glass in years. The
ultras are great from end-to-end. You also get great customer support
from both the US and Swiss offices.
Lee Cohen-Gould, M.S., C.E.M.T.
Chair, MSA Certification Board

Sr. Staff Associate in Biochemistry and
Cell & Developmental Biology
Director, Electron Microscopy & Histology and Optical Microscopy Core
Facilities Weill Cornell Medical College

lcgould-at-med.cornell.edu
voice (212)746-6146
fax (212)746-8175
http://www.med.cornell.edu/research/rea_sup/
http://www.cornellbiochem.org
http://www.cornellcelldevbiology.org

On Feb 8, 2012, at 4:46 PM, microscopylistserver-noreply-at-microscopy.com
wrote:

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Email: slakmon-at-yahoo.com Name: Slakmon

Title-Subject: [Filtered] Diamond knives & cleaning methods

Message: Good Afternoon,

I was hoping to get people's feedback and opinion on the different
diamond knives on the market. Your opinion on new, resharpened would be
helpful as well. What methods that you found best suited in cleaning
your knives without damaging your knives. Please feel free to respond on
the group
or offline directly to my email address.

Thank you in advance,

Philip

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In reply to a request to list our comments on the Listserv:

I replied to Philip but the take home message was that I do not like Blue Boats! I have used Diatome knives and the knife itself is fine and currently use their Cryo-knife.

Several of the knives that I needed to get sharpened I had sent out to Micro Star Technologies in Texas. Since I had already used their knives and liked them and their service very well I knew that they would remount the knives that I had sent in into one of their Black Boats at no additional cost. Up until last year their price was less than Diatome for sharpening. Now the two companies are close together in price as far as I remember. Both have discounts for multiple knives etc.

I was "raised" on the old DuPont knives which I loved but back in the early '80s when Diatome and DuPont (now DDK I think) would no longer make a knife below 45 degrees and had a six month waiting period, Micro Star would make any angle I wanted which was 40 degrees to cut transparent sections and it was delivered in a few weeks. Diatome does have a 35 now. I have not looked at other companies besides Micro Star Technologies to see if they also have the smaller angle knives available.

Remember to include Philip in your replies about the knives for he is not on the Listserv.
slakmon-at-yahoo.com

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E - Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}

Opinions and experiences related are those of Pat Connelly and do not represent the NIH. This message is not confidential and can be freely shared and reproduced.

________________________________
X-from: {microscopylistserver-noreply-at-microscopy.com}
Reply-To: {microscopylistserver-noreply-at-microscopy.com}

I will not reiterate too much of what others have contributed in
this discussion (most recently, Dr. Muss gave a good accounting) but
offer some personal experience.

Over the years, I've used hand-broken glass from a variety of sources,
machine-made glass knives (mostly on the LKB knifemakers), sapphire
knives (Sapphotome brand), and diamond knives from various
manufacturers (Dehmer, DuPont, Delaware Diamond Knives, Diatome). In
fact, I still have an unopened DuPont knife presented to me upon
graduation many years ago. They were all very capable knives, but I
must admit to really loving the Diatomes due to the consistently high
quality and technical support base.

Diamond knives clearly are both cost effective (if cared for properly)
and capable of cutting just about anything one would need to examine
in the TEM. Frankly, it takes less skill, as an ultramicrotomist, to
obtain high quality sections from challenging materials such as
botanicals, spores, etc. with a diamond knife. For serial sectioning,
I feel it is essential, since the replacement of glass knives during
the sectioning process causes one to lose sections.

It is much easier to obtain really thin sections with a diamond versus
glass. However, glass still has a place in the process, since it is
very useful to "face" the cutting block prior to sectioning with
diamond. Glass should always be used prior to the diamond if one is
uncertain of the nature of the specimen (to avoid damaging the
diamond). I've seen several diamonds damaged by sectioning specimens
that contained sand or glass chips.

If at all possible, only one person should be using a diamond knife,
since multiple users exponentially increase the probability of damage
to the edge. Students using diamond knives? Well, maybe really, really
good students or members of the DuPont family (i.e., rich). Good
records (maps) must be kept of the suitability of the knife edge, so
one knows which parts are free of nicks.

Cleaning of the edge will be minimized if the knife is rinsed
immediately after sectioning. Never allow the knife edge to dry
completely during any sectioning breaks. Instead, overfill the trough
so that you have a huge bulge of water and check on the knife to
verify that the edge is wet. Immediately after sectioning, I run the
section-positioning, single eyelash probe along the edge of the knife
to sweep away any debris. This single eyelash is gentle and usually
removes any debris from sections. If any remaining material is
detected (using a high magnification, well-lighted) stereomicroscope,
then some styrofoam (usually included with Diatome knives) can be
gently swept along the edge. I've had a lot of success using pith
sticks from elder (Sambucus nigra) that are touched to saliva (I know,
I know). The saliva seems to provide some lubrication and helps wet
the edge. Styrofoam, however, is safest. I've never had to resort to
using any solvents or specially designed, sonicator cleaners.

Truly, "Diamonds are a Microtomist's Best Friend," and "Diamonds are
Forever" (well, almost AND if you are very careful.

--
John J. Bozzola, Ph.D., Thankfully Retired
Professor & Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

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10, 21 -- Subject: Re: [Microscopy] Re: Diamond knives & cleaning methods
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Good Morning All

I have been following this discussion with very keen interest and have decided the Australian experience should probably also be shared.

Over the last 5 years or so there have been a number of negative experiences with Diatome knives. Those affected had preferred Diatome knives to this point and have persisted with these knives despite this. Some who had also enjoyed other brands in the past considered returning to those but didn't in the end.

The problem with the knives was generally reduced durability, the knives were not lasting a year in some cases. (That is with biological speciemns) In our experience, after the knives "wore out" we relegated them cutting semi-thin sections and saw the diamond fall out on 2 occasions.

For some satisfaction was achieved when Diatome agreed to resharpen the knives. Unfortunately, the response from Diatome and agents in Australia was generally considered to be quite patronising.

For now we continue with Diatome but all that glitters is not necessarily a Diatome knife!

Kind regards
Naomi

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Email: slakmon-at-yahoo.com Name: Slakmon

Title-Subject: [Filtered] Diamond knives & cleaning methods

Message: Good Afternoon,

I was hoping to get people's feedback and opinion on the different
diamond knives on the market. Your opinion on new, resharpened would be
helpful as well. What methods that you found best suited in cleaning
your knives without damaging your knives. Please feel free to respond on
the group
or offline directly to my email address.

Thank you in advance,

Philip

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Has anyone frozen 0.01% (w/v) Poly-L-lysine (aqueous) for storage (at 0 or -70 degrees Celsius)? We will occasionally use about 40mL at a time but don't want to waste the rest.

Thank you,

Jaci

Jaclynn Lett
Senior Research Technician
Research Center for Auditory and Visual Studies Washington University School Of Medicine Saint Louis, Missouri lettj-at-ent.wustl.edu

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The New York Society of Experimental Microscopists &
The New York Microscopical Society
Annual Joint Meeting
Wednesday February 15, 2012
5:30 PM

Weill Cornell Medical College
Room E-115 (Biochemistry)
1300 York Avenue entrance at 69th Street

Erik L. Snapp, Ph.D.
Assistant Professor
Department of Anatomy & Structural Biology
Albert Einstein College of Medicine

"Fluorescence Microscopy Applications for Cell Biology:
Practical issues with Fluorescent Proteins and measuring protein mobility in Cells"

Dr. Snapp's lab focuses on understanding the cell biology of secretory protein synthesis through a combination of live cell imaging, biophysical fluorescence methods, biochemistry, and molecular biology. His group is investigating the regulation, organization and dynamics of secretory protein translocation and folding in cells. Dr. Snapp�s talk will focus on the use of cutting edge quantitative fluorescence microscopy methods including FRAP, FLIP, photoactivation and FRET to reveal information about protein mobility in living cells.

MEETINGS ARE FREE AND OPEN TO ALL!

Frank Macaluso
Administrative Director and
Director of Electron Microscopy
Analytical Imaging Facility
Albert Einstein College of Medicine
1300 Morris Park Avenue
Bronx, NY 10461

(718) 430-3547
frank.macaluso-at-einstein.yu.edu
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Colleagues,

The Early Registration deadline for Histochemistry 2012 is Friday,
February 17. Histochemistry2012 is The Histochemical Society's Annual
Meeting and Short Course in Immunohistochemistry and Microscopy. The
Course dates are March 18-20 with the Meeting following on March 21-23.

Both meeting and course will be at MBL in Woods Hole, MA. Registration
covers all fees including accommodations and meals.

Highlights
JHC Plenary Lecture - Richard Hynes of MIT and HHMI.
Chroma Technology Lecture - Bernd Fritzsch, Univ. of Iowa
Three days and evening plenary sessions
Industry Technology Session
HCS Awards

HCS meeting information: http://www.histochemistry2012.org/
HCs registration website:
http://www.regonline.com/Register/Checkin.aspx?EventID=1011701
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HCS: http://histochemicalsociety.org/

Meg McGough
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I would like to respond to Naomi's article relating to Diatome knives. As one of two authorised agents in Australia for Diatome I am a little concerned at the negative attitude some may form regarding the level of service provided by some agents and indeed Diatome.

The following statements are made purely with the intention of stressing the support that Diatome provide for their products and in no way is to be seen as a commercial advertisement and I will apologise now if any readers misconstrue the intent of this mail.

There are numerous factors in operation when using diamond knives and all have the potential of causing rapid damage to the edge, and when this occurs, is most often perceived by many as being the result of reduced durability of the diamond edge. I propose that this belief regarding the durability is not entirely true and I suspect that there would be many individuals more conversant with diamond than I that would support this premise.

The important factor when confronted with such issues is to document the damage to the edge and try to identify the cause. Only when this is done can we propose methods designed to minimise damage to the edge and thus increase the longevity of the knife edge. As an authorised Diatome agent for some 10 years and with over 30 years experience with Diatome knives, I have been involved in many issues relating to the performance of diamond knives and can confirm that the response from Diatome over this period has been nothing short of exemplary. Diatome treat each and every complaint seriously, and in all instances which have been brought to my attention over this period of time, the knives and blocks were returned to Diatome and were extensively evaluated to determine and document the extent of the damage and possible cause. The results were provided to each user and proposals/suggestions were offered in order to help minimise future damage which could occur. This action, in most cases for those that adopted the recommendations, there was a satisfactory resolution to the issue. All knives were re-sharpened free of charge as a matter of course and good faith and returned to the customer at no cost.

I would urge any Diatome user in Australia experiencing problems with their knives, to bring it to my attention immediately and I will guarantee that we will investigate the issue promptly and make every effort to resolve the issues satisfactorily. Helmut Gnaegi from Diatome is well known throughout the International EM community and will ensure that the greatest level of support will be provided to their customers.

Graham Tranter
Emgrid Australia Pty Ltd
PO Box 118
The Patch Vic 3792
Australia
Phone: +61 3 9752 1785
Fax: +61 3 9752 1784
Mobile: 0400 060 859
Email: graham-at-emgrid.com.au
Web: www.emgrid.com.au

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Dear Listers,
I too, have to share some experience from using Diatome knives. Observing the discussion, I was hesitating whether to tell my (or better, our) experience, but seeing Naomi's comments, I thought I will share the history of Diatome knives in Seville, Spain.

Namely, a couple years ago I (our lab) bought a Diatome diamond knife for cutting ultra-thin sections of biological embedded sections. Right after seeing the first several ultras I cut, I thought "What the heck, I damaged the new knife!". All of the sections had scratch marks on them! Well, it was just opened, although bought several months prior to first use, and thinking I was the one that damaged the edge, I didn't contact the provider nor Diatome. To be clear, I consider myself an experienced user of diamond knives - I had an excellent training and very experienced teachers who taught me everything they knew, and I've been cutting for some 6 years now (not very long compared to some people on the list - hats off to them). My first thought was that I had done some very erratic thing that went unnoticed. There are only three things that touch the edge of my knife, ever: soft biological specimen embedded in EPON, cleaning stub with ethanol and occasionally an eye lash, so something might have slipped by. Well, whatever... I went on using my damaged knife.
But later, a new person came in a lab next door, and she also started with EM of biological samples. Wishing to be independent and obeying the rule of sharing (you don't share your car and your diamond), she bought a Diatome knife, because, you know, they are the best! Well, when she opened it, it looked little bit strange -the diamond edge looked little bit tilted to one side. Nothing, she went on cutting, because we thought that it might be just an optical illusion. And then, the first ultra sections. Can you guess what happened? Yep! You're right, scratches! On the very first ultra sections! Unlike me, however, she contacted Diatome trough the provider, but got a not "pleasant" response, like, (free citation) "Diatome is the best, no way the diamond can be tilted, the scratch marks are done by you, and if you want it repaired, you have to pay" (I will ask her again for the original message if one wants to see it). Negotiations between them led to a reduction in re-sharpening price. And then, after some weeks of waiting, she received her knife back, re-sharpened. And look, a miracle! The edge was not tilted anymore! (Of course that it was re-sharpened, no objection to that). And as a present for being a customer of Diatome, she received a, watch this, full box of 10 cleaning stubs! Wow, man, I'd like to receive such present, maybe I'll send my knife too!
Speaking in front of the ultramicrotome and criticizing Diatome, another person joined the communication, saying, "You have problems with your Diatome knife? Well, me too, you know, I just bought a 1.5mm diamond knife and now I can use some 0.75mm of it".
Wow, three persons, with new knives from Diatome, having the same symptoms? And all of them damaged the knife by themselves? During the first use? Well, a coincidence, you know...
And then, the technician of the core facility where we do all the EM came saying: "Don't tell me about Diatome, we've had some unpleasant experience with them". Unfortunately, we had to end the conversation at that point because of some experiments and I didn't get what kind of experience they had with Diatome.
So, four persons, or three with knives, have something to say about Diatome? Hmm, if that's not a fish, then something's fishy here...

For some time now I had an intent to ask the list about their experience with Diatome and maybe gather other people that had problems with their knives. Now, seeing this post, I used the possibility to express my thoughts and feelings regarding the subject. Sorry for the irony used, but to buy a tool of 2000€ and then be able to use just some 50% of its potential is really ·-at-#-at-|-at-][\| (frustrating might be the right word). In addition I am angry at myself for not realising the idea of recording in a video every step that I have done with the knife. At the beginning I thought it might be exaggerating, but now I realise that I made a mistake.

So, please share again your experience and if there are any other users with similar experience, please send a message, on or off line. If there is enough critical mass, maybe we could contact Diatome and do something about it?

Happy cutting,

Dr. Josif Mircheski
_______________________________________________________________________________
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/Nº
41013 Sevilla
Spain

Phone:+34-955923045
e-mail: jmircheski-at-us.es
web: www.ibis-sevilla.es

-----Original Message-----
X-from: Naomi_McCallum-at-health.qld.gov.au [mailto:Naomi_McCallum-at-health.qld.gov.au]
Sent: Friday, February 10, 2012 1:37 AM
To: jmircheski-at-us.es

Good Morning All

I have been following this discussion with very keen interest and have decided the Australian experience should probably also be shared.

Over the last 5 years or so there have been a number of negative experiences with Diatome knives. Those affected had preferred Diatome knives to this point and have persisted with these knives despite this. Some who had also enjoyed other brands in the past considered returning to those but didn't in the end.

The problem with the knives was generally reduced durability, the knives were not lasting a year in some cases. (That is with biological speciemns) In our experience, after the knives "wore out" we relegated them cutting semi-thin sections and saw the diamond fall out on 2 occasions.

For some satisfaction was achieved when Diatome agreed to resharpen the knives. Unfortunately, the response from Diatome and agents in Australia was generally considered to be quite patronising.

For now we continue with Diatome but all that glitters is not necessarily a Diatome knife!

Kind regards
Naomi

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Email: slakmon-at-yahoo.com Name: Slakmon

Title-Subject: [Filtered] Diamond knives & cleaning methods

Message: Good Afternoon,

I was hoping to get people's feedback and opinion on the different
diamond knives on the market. Your opinion on new, resharpened would be
helpful as well. What methods that you found best suited in cleaning
your knives without damaging your knives. Please feel free to respond on
the group
or offline directly to my email address.

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Philip

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Dear Listers,

Could someone share an efficient protocol for preparing mouse nerves for
TEM? I am currently using 2.5% glutaraldehyde/1% PFA in PBS (standard
fixative that I use for EM) o/n at 4� an then standard processing, but I've
encountered some problems. There are plenty of precipitate artefacts (osmium
or uranyl) between the myelin sheets and also some samples show poor resin
infiltration of the myelin. The poorly infiltrated parts of the myelin
appear to be these myelin incisures called also Schmidt-Lanterman clefts (I
thought these were processing artefacts when I first saw them).
One way to avoid these problems would be to increase time of washing and
infiltration. However, does anyone have a better protocol, that is already
successfully applied to nerves? If so, would you share it, please? Or your
comments?

Best,

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

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Josif,

I have several suggestions for you.

1. I really would not use PBS but a phosphate buffer like Millonig's or
Sorensen's at a 0.1M final concentration or I have seen quite good images
with a 0.12M Sodium Cacodylate at 7.3 pH. PBS does nasty things to cell
ultrastructure depending on how long they are exposed to it. IF you need to
wash the tissue before fixation use the above buffers.

2. There is a need with dense structures (as well as some mitochondria and
bacteria) to extend the buffer washing times between the primary and
secondary fixations. An hour with at least 3 changes would not be a bad
idea. I also give a half hour or more to washes (1 buffer and 2 water)
between the Osmium and 1% UA.

3. I assume you go through several Propylene oxide : Epon steps. These
should not be shorter than a half hour but an hour each may help.

4. Check the size of your samples. Thin ones have better infiltration so if
you have been cutting 1mm cubic pieces, cut some in half to 1x1x0.5 or
smaller.

5. Do you have a vacuum oven? You can pull a mild vacuum while the samples
are in a good volume of Epon still in a vial or I have drawn a vacuum on a
vial by placing tubing hooked to a vacuum source like an old mechanical pump
against the lid of a scintillation vial that had a small hole drilled in the
center. Let the bubbles form then release the vacuum several times rather
than drawing a strong vacuum for a longer period of time.

6. Infiltration over night with a change of Epon the next day is a common
practice. Some have said that one can put the embeddings into a cooler oven
to start, 35 to 40 degrees C. ,then after several hours put the temperature
up to 60 is helpful. This makes the Epon less thick for a longer period of
time before it starts to harden.

7. Check out unstained sections or ones only stained with lead If you are
getting too much stain on the myelin. Especially if you used the UA block
stain overnight.

8. You most likely do this but...Always make up the Ethanol dilutions from
200 proof/100% stock. The 95% alcohol that is much less expensive and used
for LM histology is distilled differently I believe and I do not understand
it but the ultrastructure is not as nice if this is used.

Wow, that was a lot longer than I intended. Hopefully you can make use of
at least one suggestion to improve your results. Frustration on not getting
good images after putting so much effort into our work is one of the things
that is so "frustrating"!

Let us know what you have done when you get the results that you are looking
for.

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E - Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}

Opinions and experiences related are those of Pat Connelly and do not
represent the NIH. This message is not confidential and can be freely shared
and reproduced.

________________________________
X-from: {jmircheski-at-us.es}
Reply-To: {jmircheski-at-us.es}

Dear All:

Early in my career, I used diamond knives of different brands, but
settled on Diatome knives because, to me, it functioned so much better
than any other knives I used. Blue color did not bother me at all. I
actually likes it better because it is unique. At one point, I had a
Diatome knife that I used for over 10 years (on a routine basis) without
resharpening and it worked just fine. To the date, I have not had any
problem with Diatome knives in my hands. However, I did notice that when
two of our new staff members used diamond knives, they were not so long
lasting. Thank you.

Hong

On 2/15/12 6:17 AM, "jmircheski-at-us.es" {jmircheski-at-us.es} wrote:

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Dear All,
I am looking for a functioning Zeiss EM 10 A/ B high voltage cable or a tip of someone repairing it or doing it new.

Best wishes,
Stefan

-------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D-97072 Wuerzburg Germany
Tel. +49-931-7848700
Fax +49-931-7848701
Mobile +49-175-7177051

Email: diller-at-stefan-diller.com
Websites: www.stefan-diller.com
www.elektronenmikroskopie.info
www. assisi.de
--------------------------------------------

==============================Original Headers==============================
4, 22 -- From stefan.diller-at-t-online.de Thu Feb 16 03:15:48 2012
4, 22 -- Received: from mailout07.t-online.de (mailout07.t-online.de [194.25.134.83])
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4, 22 -- Subject: Zeiss EM 10 A/ B high voltage cable needed
4, 22 -- From: Stefan Diller {stefan.diller-at-t-online.de}
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Stefan

I was always told that HT cables were extremely difficult to repair and the risk of a repaired cable failing or not functioning properly would be high.

Many years ago we had to cut an HT cable to remove an old microscope we were getting for spares from another lab. About a month later the HT cable on our Siemens IA broke down spectacularly but we were told by the engineers that the cut cable was of no use whatsoever and we had to shop around for a second hand one -Lukily everybody in those days had a Siemens so it only took a couple of months to source one. I hope the prognosis for a Zeiss 10 cable is better.

A lot would depend on the nature of the cable damage but if there was a break or fault halfway along the cable it's probably not even worth thinking about.

Good luck

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: stefan.diller-at-t-online.de [stefan.diller-at-t-online.de]
Sent: 16 February 2012 09:28
To: Malcolm Haswell

Dear All,
I am looking for a functioning Zeiss EM 10 A/ B high voltage cable or a tip of someone repairing it or doing it new.

Best wishes,
Stefan

-------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D-97072 Wuerzburg Germany
Tel. +49-931-7848700
Fax +49-931-7848701
Mobile +49-175-7177051

Email: diller-at-stefan-diller.com
Websites: www.stefan-diller.com
www.elektronenmikroskopie.info
www. assisi.de
--------------------------------------------

==============================Original Headers==============================
4, 22 -- From stefan.diller-at-t-online.de Thu Feb 16 03:15:48 2012
4, 22 -- Received: from mailout07.t-online.de (mailout07.t-online.de [194.25.134.83])
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15, 37 -- From malcolm.haswell-at-sunderland.ac.uk Thu Feb 16 05:31:51 2012
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15, 37 -- 11:31:44 +0000
15, 37 -- From: Malcolm Haswell {malcolm.haswell-at-sunderland.ac.uk}
15, 37 -- To: "stefan.diller-at-t-online.de" {stefan.diller-at-t-online.de} ,
15, 37 -- "Microscopy MSA
15, 37 -- (Microscopy-at-microscopy.com)" {Microscopy-at-microscopy.com}
15, 37 -- Subject: RE: [Microscopy] Zeiss EM 10 A/ B high voltage cable needed
15, 37 -- Thread-Topic: [Microscopy] Zeiss EM 10 A/ B high voltage cable needed
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Hi

Having to repair microscopes in a world where going back to the manufacturer
was the complicated method other routes were forced upon me. I always
traced organisations like medical or general high voltage companies who,
with the cable ends supplied, always did a very good job at a fraction of
the cost from the original equipment manufacturer.

I am not sure in this era of "oh we must buy from the manufacturer" if my
route is still available; my guess it is!

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide
Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: malcolm.haswell-at-sunderland.ac.uk
[mailto:malcolm.haswell-at-sunderland.ac.uk]
Sent: 16 February 2012 11:33
To: protrain-at-emcourses.com

Stefan

I was always told that HT cables were extremely difficult to repair and the
risk of a repaired cable failing or not functioning properly would be high.

Many years ago we had to cut an HT cable to remove an old microscope we were
getting for spares from another lab. About a month later the HT cable on our
Siemens IA broke down spectacularly but we were told by the engineers that
the cut cable was of no use whatsoever and we had to shop around for a
second hand one -Lukily everybody in those days had a Siemens so it only
took a couple of months to source one. I hope the prognosis for a Zeiss 10
cable is better.

A lot would depend on the nature of the cable damage but if there was a
break or fault halfway along the cable it's probably not even worth thinking
about.

Good luck

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: stefan.diller-at-t-online.de [stefan.diller-at-t-online.de]
Sent: 16 February 2012 09:28
To: Malcolm Haswell

Dear All,
I am looking for a functioning Zeiss EM 10 A/ B high voltage cable or a tip
of someone repairing it or doing it new.

Best wishes,
Stefan

-------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D-97072 Wuerzburg Germany
Tel. +49-931-7848700
Fax +49-931-7848701
Mobile +49-175-7177051

Email: diller-at-stefan-diller.com
Websites: www.stefan-diller.com
www.elektronenmikroskopie.info
www. assisi.de
--------------------------------------------

==============================Original Headers==============================
4, 22 -- From stefan.diller-at-t-online.de Thu Feb 16 03:15:48 2012
4, 22 -- Received: from mailout07.t-online.de (mailout07.t-online.de
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16 Feb 2012
15, 37 -- 11:31:44 +0000
15, 37 -- From: Malcolm Haswell {malcolm.haswell-at-sunderland.ac.uk}
15, 37 -- To: "stefan.diller-at-t-online.de" {stefan.diller-at-t-online.de} ,
15, 37 -- "Microscopy MSA
15, 37 -- (Microscopy-at-microscopy.com)" {Microscopy-at-microscopy.com}
15, 37 -- Subject: RE: [Microscopy] Zeiss EM 10 A/ B high voltage cable
needed
15, 37 -- Thread-Topic: [Microscopy] Zeiss EM 10 A/ B high voltage cable
needed
15, 37 -- Thread-Index: AQHM7JBxN2KwB2YgtEmES71SnLSLT5Y/XpO3
15, 37 -- Date: Thu, 16 Feb 2012 11:31:43 +0000
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29, 22 -- From protrain-at-emcourses.com Thu Feb 16 06:13:23 2012
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29, 22 -- Subject: RE: [Microscopy] RE: Zeiss EM 10 A/ B high voltage cable needed
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Hi Stefan,

We custom-making 50KV cable assemblies for FIB equipment:

http://www.freudlabs.com/micrion_fib_spare_parts

and I would be more then happy to look into the possibility of repairing
your HV cable.

Alternatively - look for local companies involved in servicing X-ray
equipment. Molding HV connectors has its specifics, but certainly can be
done and people servicing X-ray equipment would be very familiar with it.

Cheers :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
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On 2/16/2012 4:17 AM, stefan.diller-at-t-online.de wrote:
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} Dear All,
} I am looking for a functioning Zeiss EM 10 A/ B high voltage cable or a tip of someone repairing it or doing it new.
}
} Best wishes,
} Stefan
}
} -------------------------------------------
} Stefan Diller - Scientific Photography
} Arndtstrasse 22
} D-97072 Wuerzburg Germany
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} Email: diller-at-stefan-diller.com
} Websites: www.stefan-diller.com
} www.elektronenmikroskopie.info
} www. assisi.de
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8, 43 -- Subject: Re: [Microscopy] Zeiss EM 10 A/ B high voltage cable needed
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Perhaps I can share my experience with our DIATOME diamond knifes too.
We have both ultra and histoknifes and are satisfied with both.
I have to cut mineral microparticles embedded in resin from time to time so I got several clear marks on the edge but I cannot blame Diatome for that.
On the contrary I was surprised that the knife resisted somehow these terrible samples.
We let Diatome resharpen the knife once�and it came back just as knew.

Regards,
Stephane

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3, 39 -- Subject: Re: [Microscopy] viaWWW:Diamond knives cleaning methods
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Hello!
Documentation of a JEOL JEM-2100F on the lens and deflector system, and of the use of the "SPOT deflector system" is wanted:
I have from time to time access to a JEOL JEM-2100F, equipped with STEM functionality, but the user manual of that machine does not provide information on all present lenses and deflectors! I am trying to educating myself in practical Electron Microscopy with the help of the book of Williams & Carter, but find myself unable to translate their graphs about typical electron optical systems to the JEM-2100F lens and deflector system. Unfortunately, an intensive internet search didn�t provide me with answers either. I will not go into details about the confusion in my head, because the ones of you who better know the lens and deflector system of the JEM-2100F for sure will already have a clear idea about what I am speaking about: missing C2 lens, a so called C3 which seems not to be the upper polepiece part of the OL (as naming would be used according to Williams & Carter), the presence of an undocumented deflector system called �SPOT� most likely situated between the GUN and the condenser system, missing in the user manual an indication of the localization of the scan coils used for STEM, black-box about mysterious JEOL-engineer-password hidden alignment settings, etc...

--} I would appreciate if someone could point me to proper literature or even provide me a full scheme of the lens and deflector system of the JEM-2100, and could help me to translate the nomenclature used by JEOL to the nomenclature used �usually� in TEM lecture books and various TEM training internet sites.

--} I especially would be grateful, if somebody would have information on how to best include the �SPOT� deflector system into the alignment of TEM and STEM and when to take advantage of this system during TEM and STEM works

According to my internet search, it could be that other JEOL models of the 2000 series (2010, 2200, etc) are built up very similar, but I am not sure about this. So, please feel free to also post (or send by e-mail) information about those machines � might be better for me than not receiving anything.
Thanks a lot!!!
Robert

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8, 19 -- From robertmschmidt-at-hotmail.com Thu Feb 16 10:07:41 2012
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8, 19 -- Subject: TEM and STEM: documentation of a JEOL JEM-2100F and its "SPOT
8, 19 -- deflector system" needed
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I have used both Diatome and MicroStar diamond knives (3 of each). All of them were of good quality.

It is a good practice to check a new diamond knife as soon as you get one. Cut a few sections of pure embedding resin with different sections of a knife (I never had any scratches with a new knife). I fully agree with a manufacturer in refusing to accept complains after several months have passed since shipping a knife.

We cut a lot of hard tissue (bone and teeth), but even with them I can cut a lot of sections with the same place of a knife. (Tooth enamel is a different story: just one specimen and knife is dull). I strongly believe that main cause that is shortening the knife life (when cutting a soft tissue) is not a number of sections cut, but hard particles embedded with a tissue. I quite often stain a block faces (after sections were cut) and look at them with SEM; also I often prepare cell cultures for both TEM and SEM observation. I have seen particles of glass, carbon steel, stainless steel, titanium, and of course, SiO2 (glass, quartz, sand) in these specimens. I have seen many other particles also, containing about anything, from Al to Sb, but I have no idea about their hardness.

I always wash a knife immediately after cutting, not letting it to dry out. I wash it under a good stream of tap water (both sides of a diamond), then rinse it in DI water, and then dry it with an air stream from a duster can. I almost never use a foam stick, just do not need it. If I do have a problem with knife (such as poor wetting, sometimes stuck section), I soak it overnight in Alconox, use a foam stick if needed, then soak it in a few changes of a DI water to remove detergent. Works good for me.

Regards,

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: jmircheski-at-us.es [mailto:jmircheski-at-us.es]
} Sent: Wednesday, February 15, 2012 5:16 AM
} To: Dusevich, Vladimir
} Subject: [Microscopy] Diamond knives: Spanish experience
}
}
}
}
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} Dear Listers,
} I too, have to share some experience from using Diatome knives.
} Observing the discussion, I was hesitating whether to tell my (or
} better, our) experience, but seeing Naomi's comments, I thought I will
} share the history of Diatome knives in Seville, Spain.
}
} Namely, a couple years ago I (our lab) bought a Diatome diamond knife
} for cutting ultra-thin sections of biological embedded sections. Right
} after seeing the first several ultras I cut, I thought "What the heck,
} I damaged the new knife!". All of the sections had scratch marks on
} them! Well, it was just opened, although bought several months prior to
} first use, and thinking I was the one that damaged the edge, I didn't
} contact the provider nor Diatome. To be clear, I consider myself an
} experienced user of diamond knives - I had an excellent training and
} very experienced teachers who taught me everything they knew, and I've
} been cutting for some 6 years now (not very long compared to some
} people on the list - hats off to them). My first thought was that I had
} done some very erratic thing that went unnoticed. There are only three
} things that touch the edge of my knife, ever: soft biological specimen
} embedded in EPON, cleaning stub with ethanol and occasionally an eye
} lash, so s!
} omething might have slipped by. Well, whatever... I went on using my
} damaged knife.
} But later, a new person came in a lab next door, and she also started
} with EM of biological samples. Wishing to be independent and obeying
} the rule of sharing (you don't share your car and your diamond), she
} bought a Diatome knife, because, you know, they are the best! Well,
} when she opened it, it looked little bit strange -the diamond edge
} looked little bit tilted to one side. Nothing, she went on cutting,
} because we thought that it might be just an optical illusion. And then,
} the first ultra sections. Can you guess what happened? Yep! You're
} right, scratches! On the very first ultra sections! Unlike me, however,
} she contacted Diatome trough the provider, but got a not "pleasant"
} response, like, (free citation) "Diatome is the best, no way the
} diamond can be tilted, the scratch marks are done by you, and if you
} want it repaired, you have to pay" (I will ask her again for the
} original message if one wants to see it). Negotiations between them led
} to a reduction in re-sharpen!
} ing price. And then, after some weeks of waiting, she received her
} knife back, re-sharpened. And look, a miracle! The edge was not tilted
} anymore! (Of course that it was re-sharpened, no objection to that).
} And as a present for being a customer of Diatome, she received a, watch
} this, full box of 10 cleaning stubs! Wow, man, I'd like to receive such
} present, maybe I'll send my knife too!
} Speaking in front of the ultramicrotome and criticizing Diatome,
} another person joined the communication, saying, "You have problems
} with your Diatome knife? Well, me too, you know, I just bought a 1.5mm
} diamond knife and now I can use some 0.75mm of it".
} Wow, three persons, with new knives from Diatome, having the same
} symptoms? And all of them damaged the knife by themselves? During the
} first use? Well, a coincidence, you know...
} And then, the technician of the core facility where we do all the EM
} came saying: "Don't tell me about Diatome, we've had some unpleasant
} experience with them". Unfortunately, we had to end the conversation at
} that point because of some experiments and I didn't get what kind of
} experience they had with Diatome.
} So, four persons, or three with knives, have something to say about
} Diatome? Hmm, if that's not a fish, then something's fishy here...
}
} For some time now I had an intent to ask the list about their
} experience with Diatome and maybe gather other people that had problems
} with their knives. Now, seeing this post, I used the possibility to
} express my thoughts and feelings regarding the subject. Sorry for the
} irony used, but to buy a tool of 2000�,� and then be able to use just
} some 50% of its potential is really ·-at-#-at-|-at-][\| (frustrating might be
} the right word). In addition I am angry at myself for not realising the
} idea of recording in a video every step that I have done with the
} knife. At the beginning I thought it might be exaggerating, but now I
} realise that I made a mistake.
}
} So, please share again your experience and if there are any other users
} with similar experience, please send a message, on or off line. If
} there is enough critical mass, maybe we could contact Diatome and do
} something about it?
}
} Happy cutting,
}
} Dr. Josif Mircheski
} _______________________________________________________________________
} ________
} Instituto de Biomedicina de Sevilla (IBIS), Lab 108 Campus del Hospital
} Universitario Virgen del Rocio/CSIC/Univ. de Sevilla Avda. Manuel
} Siurot S/Nº
} 41013 Sevilla
} Spain
}
} Phone:+34-955923045
} e-mail: jmircheski-at-us.es
} web: www.ibis-sevilla.es
}
} -----Original Message-----
} X-from: Naomi_McCallum-at-health.qld.gov.au
} [mailto:Naomi_McCallum-at-health.qld.gov.au]
} Sent: Friday, February 10, 2012 1:37 AM
} To: jmircheski-at-us.es
} Subject: [Microscopy] Fwd: viaWWW:Diamond knives & cleaning methods
}
}
}
}
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} Good Morning All
}
} I have been following this discussion with very keen interest and have
} decided the Australian experience should probably also be shared.
}
} Over the last 5 years or so there have been a number of negative
} experiences with Diatome knives. Those affected had preferred Diatome
} knives to this point and have persisted with these knives despite this.
} Some who had also enjoyed other brands in the past considered returning
} to those but didn't in the end.
}
} The problem with the knives was generally reduced durability, the
} knives were not lasting a year in some cases. (That is with biological
} speciemns) In our experience, after the knives "wore out" we relegated
} them cutting semi-thin sections and saw the diamond fall out on 2
} occasions.
}
} For some satisfaction was achieved when Diatome agreed to resharpen the
} knives. Unfortunately, the response from Diatome and agents in
} Australia was generally considered to be quite patronising.
}
} For now we continue with Diatome but all that glitters is not
} necessarily a Diatome knife!
}
} Kind regards
} Naomi
}
} } } } {microscopylistserver-noreply-at-microscopy.com} 2/9/2012 7:45 am } } }
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} Title-Subject: [Filtered] Diamond knives & cleaning methods
}
} Message: Good Afternoon,
}
} I was hoping to get people's feedback and opinion on the different
} diamond knives on the market. Your opinion on new, resharpened would be
} helpful as well. What methods that you found best suited in cleaning
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Robert;
The missing C2 in JEOL FEG microscopes is indeed confusing without a little history: The 2010 LaB6 microscope had three condenser lenses. The 2010F did not need three because less demagnification was needed for a FEG source. The middle lens, or C2 was taken out, and all the JEOL FEGs I have seen (2010F, 2100F and ARM 200) use a nomenclature of C1 and C3 for the two condenser lenses.

John Mardinly, ASU

-----Original Message-----
X-from: robertmschmidt-at-hotmail.com [mailto:robertmschmidt-at-hotmail.com]
Sent: Thursday, February 16, 2012 9:14 AM
To: John Mardinly

Hello!
Documentation of a JEOL JEM-2100F on the lens and deflector system, and of the use of the "SPOT deflector system" is wanted:
I have from time to time access to a JEOL JEM-2100F, equipped with STEM functionality, but the user manual of that machine does not provide information on all present lenses and deflectors! I am trying to educating myself in practical Electron Microscopy with the help of the book of Williams & Carter, but find myself unable to translate their graphs about typical electron optical systems to the JEM-2100F lens and deflector system. Unfortunately, an intensive internet search didn�t provide me with answers either. I will not go into details about the confusion in my head, because the ones of you who better know the lens and deflector system of the JEM-2100F for sure will already have a clear idea about what I am speaking about: missing C2 lens, a so called C3 which seems not to be the upper polepiece part of the OL (as naming would be used according to Williams & Carter), the presence of an undocumented deflector system called "SPOT" most likely situated between the GUN and th!

e condenser system, missing in the user manual an indication of the localization of the scan coils used for STEM, black-box about mysterious JEOL-engineer-password hidden alignment settings, etc...

--} I would appreciate if someone could point me to proper literature or even provide me a full scheme of the lens and deflector system of the JEM-2100, and could help me to translate the nomenclature used by JEOL to the nomenclature used "usually" in TEM lecture books and various TEM training internet sites.

--} I especially would be grateful, if somebody would have information on how to best include the "SPOT" deflector system into the alignment of TEM and STEM and when to take advantage of this system during TEM and STEM works

According to my internet search, it could be that other JEOL models of the 2000 series (2010, 2200, etc) are built up very similar, but I am not sure about this. So, please feel free to also post (or send by e-mail) information about those machines - might be better for me than not receiving anything.
Thanks a lot!!!
Robert

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Robert

There is a cross sectional diagram of the microscope in the JEOL
instructions which shows the location of all lenses and alignments
necessary for operation.

John has already summarized the reason for labelling the two condenser
lenses C1, C3. For field emission less demag is needed so there is only one
condenser lens (C1) for LaB6 systems more demag is needed to reduce the
spot size so two lenses are used (C1,& C2). In both cases C3 is used to
spread the beam on the screen (Brightness) in TEM mode. Some JEOL field
emission TEM drawings will show C1, C3 some C1, C2! The condenser lens in
the upper bore of the objective lens is the condenser mini-lens (CM) and is
part of the C/O lens design. By adding the CM lens directly above the
objective we can introduce a crossover in front of the objective that will
result in a parallel illumination at the specimen for TEM while retaining
good probe forming capabilities when it is off (STEM).

SPOT align is an engineer level alignment setting used to get the shift
alignment approximately correct as you change probe sizes. I believe that
they are part of the condenser stigmator assembly just before the condenser
(beam alignment) but after C3 (C2). They should not be used in normal
operation. Movements due to changing probe size are corrected using gun
shifts and condenser (beam) shifts.

STEM scanning is done by scanning the condenser alignment coils.

Hope this helps.

Alan

At 10:08 AM 2/16/2012, you wrote:

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Good Morning Everyone,

Does anyone on the list have any experience in etching nylon with concentrated formic acid? I have a procedure, but the MSD sheet describes formic acid as reactive with water. I suspect they mean reactive as are all concentrated acids are with water. But it got me thinking, What do I do with the small amount (total 15 ml) of acid after etch?

I'd like to dilute and dispose, but....

Any suggestions on etching and disposal would be very welcome.

Thanks!

Frank Karl
Microscopist
ARDL
2887 Gilchrist Road
Akron, Ohio 44305
330-794-6600

________________________________
This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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I can't resist (it is Friday, after all!)

Since formic acid is an ant-acid, shouldn't it be able to neutralize
itself? ;-)

JME

--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

My grandfather once told me that there are
two kinds of people: those who work and those
who take the credit. He told me to try to be
in the first group; there was less competition there.
- Indira Gandhi

On 17/02/2012 9:28 AM, frank_karl-at-ardl.com wrote:
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} Good Morning Everyone,
}
} Does anyone on the list have any experience in etching nylon with concentrated formic acid? I have a procedure, but the MSD sheet describes formic acid as reactive with water. I suspect they mean reactive as are all concentrated acids are with water. But it got me thinking, What do I do with the small amount (total 15 ml) of acid after etch?
}
} I'd like to dilute and dispose, but....
}
} Any suggestions on etching and disposal would be very welcome.
}
}
} Thanks!
}
} Frank Karl
} Microscopist
} ARDL
} 2887 Gilchrist Road
} Akron, Ohio 44305
} 330-794-6600
}
}
}
} ________________________________
} This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.
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Just use plenty of water.
It's an acid even the body can produce and breakdown.
It's part of the alcohol breakdown cycle.
So I would not be to concerned.
Just safety goggles and plenty water
My 2 cents

--
With kind regards,
Gert ten Brink
Dept. Nanostructured Materials and Interfaces
Zernike Institute for Advanced Materials
University of Groningen
Nijenborgh 4
9747 AG Groningen
The Netherlands
phone: +31(0)503634889; fax:+31(0)503634881
e-mail: g.h.ten.brink-at-rug.nl;
http://www.m2i.nl

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Dear Frank,

right now I was working on a compilation for the Disposal of Formic Acid for our Hospital...so:
as with the former posters..:

it might be dangerous to work with undiluted formic acid without "respect"... so you should work in/under a fume hood and wear perhaps gloves and safety goggles (just to be on the very safe side: do not inhale!).

After the reaction:

here in Europe we it would be desirable also to dispose of formic acid as a {special and dangerous} waste, but only if it were in Liters.
For the small amount of 15-100 ml you use (if you don't mind that I increased the volume a little bit):

just dilute carefully and thoroughly with plenty of only plain water
(and if you would like to play with it, you also in addition can neutralize with anorganic bases like NaOH)
before you spill it down the sink.

Just for fun: bees, ants, stinging nettle and even fir needles (yes, and tobacco smoke) contain formic acid...

Best wishes and regards,
have a nice and beautiful weekend,

Wolfgang Muss
SALZBURG, Austria

} -----Urspr�ngliche Nachricht-----
} Von: frank_karl-at-ardl.com [mailto:frank_karl-at-ardl.com]
} Gesendet: Freitag, 17. Februar 2012 14:32
} An: Mu� Wolfgang
} Betreff: [Microscopy] Formic acid etch and disposal
}
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} Good Morning Everyone,
}
} Does anyone on the list have any experience in etching nylon with
} concentrated formic acid?
}
} I have a procedure, but the MSD sheet describes formic acid as reactive
} with water.
}
} I suspect they mean reactive as are all concentrated acids are with
} water.
}
} But it got me thinking, What do I do with the small amount (total 15
} ml) of acid after etch?
}
} I'd like to dilute and dispose, but....
}
} Any suggestions on etching and disposal would be very welcome.
}
}
} Thanks!
}
} Frank Karl
} Microscopist
} ARDL
} 2887 Gilchrist Road
} Akron, Ohio 44305
} 330-794-6600
}
}
}
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Brought to you through the wonders of eBay:

http://www.ebay.com/itm/Educational-Insights-Geosafari-Jr-Talking-Electron-Microscope-180-images-/310349953149?pt=LH_DefaultDomain_0&hash=item48424c787d

and don't miss this one:

http://www.ebay.com/itm/USB-PC-Magnifying-Zoom-Bionic-Eye-Electron-Microscope-/270778420040?pt=LH_DefaultDomain_0&hash=item3f0ba6cb48

I just hope Administration doesn't find out about these low-cost
alternatives.

--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

My grandfather once told me that there are
two kinds of people: those who work and those
who take the credit. He told me to try to be
in the first group; there was less competition there.
- Indira Gandhi

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just a little note: many years ago during summer a colleague got a
minute spilling of concentrated (!) formic acid on the skin of his
forarm. He rinsed the area with a lot of water 2 minutes later, however,
in this area developed a really nasty eczema which was present for many
weeks after.
The lesson for me was: use diluted formic acid if possible.
( otherwise dispose of with a lot of tap water.)

greetings,
Peter Heimann

--
====================================================

Dr. Peter Heimann Raum: W7-107 / Tel.: +49-(0)521-106-5628
Universitaet Bielefeld Fax: +49-(0)521-106-5654
"Zellbiologie / Cell Biology" W7-107
33501 Bielefeld, Germany www.uni-bielefeld.de/biologie/cellbio

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Shameless spam. Shameless advertising.
Hope never see it again.

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: jehrman-at-mta.ca [mailto:jehrman-at-mta.ca]
} Sent: Friday, February 17, 2012 8:38 AM
} To: Dusevich, Vladimir
} Subject: [Microscopy] ****low**** priced electron microscopes
}
}
}
}
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} Brought to you through the wonders of eBay:
}
} http://www.ebay.com/itm/Educational-Insights-Geosafari-Jr-Talking-
} Electron-Microscope-180-images-
} /310349953149?pt=LH_DefaultDomain_0&hash=item48424c787d
}
} and don't miss this one:
}
} http://www.ebay.com/itm/USB-PC-Magnifying-Zoom-Bionic-Eye-Electron-
} Microscope-/270778420040?pt=LH_DefaultDomain_0&hash=item3f0ba6cb48
}
} I just hope Administration doesn't find out about these low-cost
} alternatives.
}
} --
}
} James M. Ehrman
} Digital Microscopy Facility
} Mount Allison University
} 63B York St.
} Sackville, NB E4L 1G7
} CANADA
}
} phone: 506-364-2519
} fax: 506-364-2505
} email: jehrman-at-mta.ca
} www: http://www.mta.ca/dmf
}
} My grandfather once told me that there are two kinds of people: those
} who work and those who take the credit. He told me to try to be in the
} first group; there was less competition there.
} - Indira Gandhi
}
}
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Dear Robert, Dear all!
I followed this thread with great interest, because I am this days coincidentally with our JEOL service engineer discussing almost the identical topic as you brought it up here on the list server. I agree with you about the unsatisfying quality of the manual available to the customer. It helps the beginner to quickly get up and running, but for a more thorough insight into the 2100F it is not sufficiently well informing. More in-depths points stay cryptic. I have seen even two different versions of the user manual, and from one manual to the other different ray diagrams about the usage of lenses in the various EM modes are provided (i.e. having CM in use or switched off for NBD mode)! If you really want to know how your 2100F works, better monitor the LENS and DEFLECTOR values yourself. The JEOL service engineers obviously have more complete (and hopefully more reliable) written information and more in-depth trainings from JEOL available, but unfortunately it seems as if this information has to be kept confidential by them.

Robert, to share your confusion, but also trying to answer some part of your question: there are two “classical” condenser lenses present in that TEM. The 1st one is called CL1 and the 2nd one is called CL3 for reasons which John and Alan clarified already. The 2nd condenser lens is called CL2 in the manual´s chapter 2.2.2, but it is called CL3 in the manual´s chapter 3.5.1 and in the TEMCON software. It can be controlled by the “Brightness”-Wheel of the left panel. It acts as the CL2 as explained in a classical TEM lecture (for instance in your Williams and Carter book). The manual´s chapter 2.2.2 specifies “Condenser Lens system: 3-stage (1st CL1, 2nd CL2, and condenser mini lens CM)”. Thus, in addition to the CL1 and CL3, also the CM should be treated as part of the illumination (condenser) system and not as part of the condenser-objective lens system. This is also reflected like this in the ray diagrams in chapter 3.5.1 .
Well, here easily arises confusion: as Alan already pointed out, and if you study the drawings 3.3.2 and 3.3.3 carefully, and if we add assumptions from just studying the outside of the column of the 2100F, you can see that the CM of this microscope most likely is the upper pole piece coil of a so called "condenser-objective lens" system. This is contradicting to the graphs in chapter 3.5.1 and the statement in chapter 2.2.2, though.

The manual then states in chapter 2.2.4 “Objective Lens system: double objective lens (objective lens and objective minilens)”, and “Focusing: digital coarse/fine controls”.
In the condenser-objective lens system of the 2100F, the lower pole piece coil appears as the OL. In agreement with the specification in chapter 2.2.4, there are in the TEMCON software in the Free Lens Control menu for the OL two values “OL coarse” and “OL fine” adjustable.
Well, here again easily arises confusion: does this mean that there are two coils in the lower pole piece region present as assumable from drawing 3.3.2, or do this two values act together on only one coil as assumable from drawings 3.3.3 and 3.5.1?

Although I am by far not a real expert on electron optics, I can´t agree completely on Alan´s comment about the CM functionality in this particular microscope. As we find only the CM alone being present in the upper pole piece region and do not find there another condensing lens nearby in front of the CM, and as we find that this microscope´s so called CL3 actually performs as a classical CL2 and is not part of the condenser-objective lens system, how is it possible to form with solely the CM either switched on in TEM mode a parallel illumination or switched off in STEM mode a convergent probe? I would expect that there has to be a 2nd lens present in the upper pole piece region, but can´t find it. If there is no 2nd lens closely above the CM in the upper pole piece region present, then there wouldn´t really exist a true condenser-objective lens system in this microscope, but it would just be a traditional objective lens with two separately operated pole pieces. But this could (in my opinion) not give us the excellent performance of this microscope to be able to switch so easily and nicely to both, parallel or convergent probe illumination.
Robert, I guess that we share confusion here, and hopefully the experts can teach us on this, after you gratefully brought this topic up on the list server.

About the SPOT alignment: there are indeed no special deflector coils drawn for this in any figure of the manual. And inspecting the outside of the column you will also not find a dedicated cable labeled SPOT (or an unlabeled cable) going to the column. I have to add, that at least on our microscope all cables are nicely labeled. Although the SPOT alignment deflectors have in the deflector value monitor unique values and changing this values seems to not manipulate the GUNA or CLA shift/tilt values, I expect that the SPOT deflection is a virtual function acting concerted so little on GUNA and CLA, that we don´t easily see a change in those values (GUNA and CLA) upon changing the (virtual) SPOT values. This would also explain why there is only one X and Y value pair listed, but not two pairs as usually needed for a nice beam shift proceeded by a double reflector system with a pair of coils (as you find them for instance for the GUNA and CLA). However, I picked up from our JEOL service engineer, that when operating in alpha-1 mode the SPOT deflector function should be used for centering the beam on the screen, while the SHIFTs (CLA) should stay untouched then. I also picked up from a JEOL STEM training powerpoint presentation published somewhere in the internet, that re-centering the beam in STEM mode after changing the spotsize (CL1) should be done with the SPOT deflectors, while the SHIFTs here again should stay untouched then. But in our TEM and STEM manuals this right away is documented different, they just don´t mention to use the SPOT deflectors anywhere. You will little by little find out, that there are more things present in the 2100F, which are not documented.
As Alan already pointed out, in principal you here completely depend on your JEOL service engineer to get the preset SPOT alignment well done for you. And then you might just not need to touch this setting on your own no more. Hope you have a good engineer around...

...and hope I didn´t confuse more than I could help!
Best greetings from the EM-Labs of CIC biomaGUNE,
Marco Moller

----------------------------------------------------------------
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Platform Manager - Electron Microscopy

CIC biomaGUNE
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A talking microscope! Amazing! What's next, color?

John Mardinly, ASU

}
}
}
}
} Brought to you through the wonders of eBay:
}
} http://www.ebay.com/itm/Educational-Insights-Geosafari-Jr-Talking-Electron-Microscope-180-images-/310349953149?pt=LH_DefaultDomain_0&hash=item48424c787d
}
} and don't miss this one:
}
} http://www.ebay.com/itm/USB-PC-Magnifying-Zoom-Bionic-Eye-Electron-Microscope-/270778420040?pt=LH_DefaultDomain_0&hash=item3f0ba6cb48
}
} I just hope Administration doesn't find out about these low-cost
} alternatives.
}
} --
}
} James M. Ehrman
} Digital Microscopy Facility
} Mount Allison University
} 63B York St.
} Sackville, NB E4L 1G7
} CANADA
}
} phone: 506-364-2519
} fax: 506-364-2505
} email: jehrman-at-mta.ca
} www: http://www.mta.ca/dmf
}
} My grandfather once told me that there are
} two kinds of people: those who work and those
} who take the credit. He told me to try to be
} in the first group; there was less competition there.
} - Indira Gandhi
}
}
} ==============================Original Headers==============================
} 10, 19 -- From jehrman-at-mta.ca Fri Feb 17 08:37:49 2012
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Ah, but you'll notice the images *are* in color! And SEM images to boot,
coming out of a TEM-looking device.

I've been doing it the hard way, I guess!

JME

--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

Give me ambiguity or give me something else.

On 17/02/2012 1:10 PM, John Mardinly wrote:
} A talking microscope! Amazing! What's next, color?
}
} John Mardinly, ASU
}
} }
} }
} }
} } Brought to you through the wonders of eBay:
} }
} } http://www.ebay.com/itm/Educational-Insights-Geosafari-Jr-Talking-Electron-Microscope-180-images-/310349953149?pt=LH_DefaultDomain_0&hash=item48424c787d
} }
} } and don't miss this one:
} }
} } http://www.ebay.com/itm/USB-PC-Magnifying-Zoom-Bionic-Eye-Electron-Microscope-/270778420040?pt=LH_DefaultDomain_0&hash=item3f0ba6cb48
} }
} } I just hope Administration doesn't find out about these low-cost
} } alternatives.
} }
} } --
} }
} } James M. Ehrman
} } Digital Microscopy Facility
} } Mount Allison University
} } 63B York St.
} } Sackville, NB E4L 1G7
} } CANADA
} }
} } phone: 506-364-2519
} } fax: 506-364-2505
} } email: jehrman-at-mta.ca
} } www: http://www.mta.ca/dmf
} }
} } My grandfather once told me that there are
} } two kinds of people: those who work and those
} } who take the credit. He told me to try to be
} } in the first group; there was less competition there.
} } - Indira Gandhi
} }
} }
} } ==============================Original Headers==============================
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--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

Give me ambiguity or give me something else.

==============================Original Headers==============================
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17, 21 -- References: {201202171444.q1HEib9H003257-at-ns.microscopy.com} {779138A8-9BF1-4B00-A802-56933531C553-at-exchange.asu.edu}
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And it isn't even a bargain. You can get the same at numerous sources for
about $40.00

Debby
--
Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy/

} From: "John.Mardinly-at-asu.edu" {John.Mardinly-at-asu.edu}
} Reply-To: "John.Mardinly-at-asu.edu" {John.Mardinly-at-asu.edu}
} Date: Fri, 17 Feb 2012 12:11:55 -0500
} To: Debby Sherman {dsherman-at-purdue.edu}
} Subject: [Microscopy] Re: ****low**** priced electron microscopes
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
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}
} A talking microscope! Amazing! What's next, color?
}
} John Mardinly, ASU
}
} }
} }
} }
} }
} } Brought to you through the wonders of eBay:
} }
} } http://www.ebay.com/itm/Educational-Insights-Geosafari-Jr-Talking-Electron-Mi
} } croscope-180-images-/310349953149?pt=LH_DefaultDomain_0&hash=item48424c787d
} }
} } and don't miss this one:
} }
} } http://www.ebay.com/itm/USB-PC-Magnifying-Zoom-Bionic-Eye-Electron-Microscope
} } -/270778420040?pt=LH_DefaultDomain_0&hash=item3f0ba6cb48
} }
} } I just hope Administration doesn't find out about these low-cost
} } alternatives.
} }
} } --
} }
} } James M. Ehrman
} } Digital Microscopy Facility
} } Mount Allison University
} } 63B York St.
} } Sackville, NB E4L 1G7
} } CANADA
} }
} } phone: 506-364-2519
} } fax: 506-364-2505
} } email: jehrman-at-mta.ca
} } www: http://www.mta.ca/dmf
} }
} } My grandfather once told me that there are
} } two kinds of people: those who work and those
} } who take the credit. He told me to try to be
} } in the first group; there was less competition there.
} } - Indira Gandhi
} }
} }
} } ==============================Original Headers==============================
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7, 28 -- From dsherman-at-purdue.edu Fri Feb 17 12:47:42 2012
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7, 28 -- From: "Sherman, Debra M" {dsherman-at-purdue.edu}
7, 28 -- To: "message to: MSA list" {microscopy-at-microscopy.com}
7, 28 -- Date: Fri, 17 Feb 2012 13:47:39 -0500
7, 28 -- Subject: Re: [Microscopy] Re: ****low**** priced electron microscopes
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7, 28 -- Thread-Index: Acztl0B8mj98ibiOQ2ugnNVdtkZSFAADV7Ts
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We can make fun of these toys, folks, but there's a serious message
here. These toys don't just appear on eBay; you'll find them in
major school supply catalogs. Parents and teachers BUY them. There
will be 3 educational outreach sessions at M&M 2012 in Phoenix.
Please come to one (or more!) and learn how YOU can help to fight
ignorance. If you won't be in Phoenix, please visit Project MICRO at
the URL below.

Caroline
--
--
Caroline Schooley
Project MICRO Coordinator
Microscopy Society of America
45301 Caspar Point Road, Box 117
Caspar, CA 95420
Phone/FAX (707)964-9460
Project MICRO: http://www.microscopy.org/education/projectMICRO

==============================Original Headers==============================
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2, 16 -- To: Microscopy-at-Microscopy.Com
2, 16 -- From: Caroline Schooley {schooley-at-mcn.org}
2, 16 -- Subject: [Microscopy] Re: ****low**** priced electron microscopes
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I apologize for my post, but it was not without silver lining for me: I have found out that I can be even dumber than I thought was possible.

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: DusevichV-at-umkc.edu [mailto:DusevichV-at-umkc.edu]
} Sent: Friday, February 17, 2012 9:03 AM
} To: Dusevich, Vladimir
} Subject: [Microscopy] RE: ****low**** priced electron microscopes
}
}
}
}
} -----------------------------------------------------------------------
} -----
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
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} -----------------------------------------------------------------------
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}
} Shameless spam. Shameless advertising.
} Hope never see it again.
}
} Vladimir M. Dusevich, Ph.D.
} Electron Microscope Lab Director
} 371 School of Dentistry
} 650 E. 25th Street
} Kansas City, MO 64108-2784
}
} } -----Original Message-----
} } From: jehrman-at-mta.ca [mailto:jehrman-at-mta.ca]
} } Sent: Friday, February 17, 2012 8:38 AM
} } To: Dusevich, Vladimir
} } Subject: [Microscopy] ****low**** priced electron microscopes
} }
} }
} }
} }
} } ---------------------------------------------------------------------
} --
} } -----
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} } America To Subscribe/Unsubscribe --
} } http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ---------------------------------------------------------------------
} --
} } -----
} }
} } Brought to you through the wonders of eBay:
} }
} } http://www.ebay.com/itm/Educational-Insights-Geosafari-Jr-Talking-
} } Electron-Microscope-180-images-
} } /310349953149?pt=LH_DefaultDomain_0&hash=item48424c787d
} }
} } and don't miss this one:
} }
} } http://www.ebay.com/itm/USB-PC-Magnifying-Zoom-Bionic-Eye-Electron-
} } Microscope-/270778420040?pt=LH_DefaultDomain_0&hash=item3f0ba6cb48
} }
} } I just hope Administration doesn't find out about these low-cost
} } alternatives.
} }
} } --
} }
} } James M. Ehrman
} } Digital Microscopy Facility
} } Mount Allison University
} } 63B York St.
} } Sackville, NB E4L 1G7
} } CANADA
} }
} } phone: 506-364-2519
} } fax: 506-364-2505
} } email: jehrman-at-mta.ca
} } www: http://www.mta.ca/dmf
} }
} } My grandfather once told me that there are two kinds of people:
} those
} } who work and those who take the credit. He told me to try to be in
} the
} } first group; there was less competition there.
} } - Indira Gandhi
} }
} }
} } ==============================Original
} } Headers==============================
} } 10, 19 -- From jehrman-at-mta.ca Fri Feb 17 08:37:49 2012 10, 19 --
} } Received: from smtpx.mta.ca (smtpx.mta.ca [138.73.1.4])
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} } Feb 2012 10:37:51 -0400 10, 19 -- From: "James M. Ehrman"
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} 4, 29 -- From: "Dusevich, Vladimir" {DusevichV-at-umkc.edu}
} 4, 29 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
} 4, 29 -- Subject: RE: [Microscopy] ****low**** priced electron
} microscopes
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} microscopes
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6, 29 -- Fri, 17 Feb 2012 12:55:31 -0600
6, 29 -- From: "Dusevich, Vladimir" {DusevichV-at-umkc.edu}
6, 29 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
6, 29 -- Subject: RE: [Microscopy] RE: ****low**** priced electron microscopes
6, 29 -- Thread-Topic: [Microscopy] RE: ****low**** priced electron microscopes
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Hi Folks,

We are in need of a standard holder for our Philips CM-10 microscope. Ones
from Philips EM-400 will work as well as ones from the CM series with manual
stages. Would appreciate you contacting me off list if you have a spare or
are planning to replace one of these model microscopes so may have a sample
holder you will not be needing.

Thanks,
Debby
--
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy/

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Colleagues,

A reminder that early registration for The Histochemical Society's Annual
Meeting and Short Course ends today.

The Registration site is here:
http://www.regonline.com/Register/Checkin.aspx?EventID=1011701.

The Course dates are March 18-20 with the Meeting following on March 21-23.

Both meeting and course will be at MBL in Woods Hole, MA. Registration
covers all fees including accommodations and meals.

Highlights
JHC Plenary Lecture - Richard Hynes of MIT and HHMI.
Chroma Technology Lecture - Bernd Fritzsch, Univ. of Iowa
Three days and evening plenary sessions
Industry Technology Session
HCS Awards

HCS meeting information: http://www.histochemistry2012.org/
HCS registration website:
http://www.regonline.com/Register/Checkin.aspx?EventID=1011701
HCS short course information: http://immunohistochem.com/
HCS: http://histochemicalsociety.org/

Thank you,
Meg McGough

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Email: Calum.Dickinson-at-gmail.com Name: Calum Dickinson

Organization: University of Limerick

Title-Subject: [Filtered] Position Available: Electron Microscopist (Postdoctoral Researcher)

Message: University of Limerick

Materials and Surface Science Institute, University of Limerick, Ireland

Electron Microscopist (Postdoctoral Researcher)

Contract Duration 5 years

Application Deadline 09 March 2012 12:00

The Materials & Surface Science Institute (www.ul.ie/mssi), at the University of Limerick, provides
a centre of excellence generating fundamental research on topics of industrial significance in the
fields of materials and surface science. The Institute houses a multidisciplinary team of scientists
and engineers, who undertake research focused on the design of materials for Health, Transport,
Energy, ICT and Clean Technology. There are five instrument scientists based in MSSI who oversee the
operation of the instruments, develop new techniques and collaborate with researchers in MSSI and in
industry. The Institute requires an Electron Microscopist to support and develop the microscopy
suite, which includes a JEOL JEM-2100F field emission TEM, with STEM attachment for bright field
(BF) and dark field (HAADF) imaging, a JEOL JEM 2011 microscope, a Hitachi SU-70 high resolution SEM
with EDS, WDS and EBSD capability and a Focused Ion Beam (FIB 200) with secondary ion mass
spectrometry.

We are seeking an outstanding candidate who is self-motivated, creative and career-oriented to join
our research team. The successful candidate will report to the Director of MSSI or the nominee of
the Director and will support the research activities of the Institute. Applicants must have a PhD
in materials science/electron microscopy/ chemistry/physics/engineering or a related discipline. The
successful candidate must be able to work individually and as part of a team and will:

· Have demonstrable knowledge of crystallography and diffraction techniques and extensive
experience of scanning and transmission electron microscopy (SEM and TEM) of a range of materials,
including nanomaterials, as evidenced by publication in peer reviewed journals.

Full details are available at http://www.ul.ie/hrvacancies/ Job ID 007547

Informal enquiries regarding the posts may be directed to:

Professor Noel ODowd

Materials and Surface Science Institute

University of Limerick

Email: noel.odowd-at-ul.ie

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Marco: A loooong answer. Thanks that you admitted where assumptions are leaving questions unresolved. Let�s see if even more answers will be posted, someone should be able to overcome remaining assumptions. Don�t get me wrong. I�m very glad for your analysis of the situation. Very helpful for me. So, THANK YOU!
Robert.

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Dear Listers,

I am in a need of a proven protocol for preparing motor end-plates (NMJ) for
SEM. I've found several that employ use of 8N HCL and then collagenase, but
I find this treatment to harsh for my gentle muscle. Moreover, these papers
show only the post-synaptic part, while I need to focus on the presynaptic
part.
Any advice would be welcomed.

As always, greetings from Seville,
Josif

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

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Good evening,
Dear Josif,

unfortunately I only do have the following paper available (and it deals - as judged from the micrographs and from the text - with subneural apparatuses (SNAs) described as elaborate postsynaptic structures particularly well developed at mammalian neuromuscular junctions)

=========================
Letter
Aged neuromuscular junctions in the extensor digitorum longus muscle of the rat as revealed
by scanning electron microscopy
Sadaaki Oki, Junzo Desaki, Taichi Ezaki, and Yoshiro Matsuda
Departments of Orthopaedic Surgery and Anatomy, Ehime University School of Medicine, Shigenobu,
Ehime 791-0295, and Department of Anatomy, Kumamoto University School of Medicine, Honjo,
Kumamoto 860-0811, Japan

Journal of Electron Microscopy 48(3): 297-300 (1999)
==========================
References (in total 17) for the method used (The specimens after thorough fixation were "treated with a HCl-hydrolysis procedure" [
Desaki J and Uehara J {1981} The overall morphology of neuromuscular junctions as revealed by Scanning Electron Microscopy])
"Find" {pre-syn} in the text: 0 result. (.pdf available if you don't have this one)

The problem will be that if you use also a moderate HCL-hydrolysis you most probably will "etch" away also the delicate nerve endings you would like to see, so perhaps long / thorough fixation and an (T)OTOTO stain-procedure followed by CPD and {microscopically} playing around with detaching layers of tissue at NMJ's could lead to a result...(unfortunately not knowing an alternative method for the pre-synaptic side of NMJ).

Sorry if this is not helpful to you,
best wishes and regards,

Wolfgang MUSS
SALZBURG-Austria

} -----Urspr�ngliche Nachricht-----
} Von: jmircheski-at-us.es [mailto:jmircheski-at-us.es]
} Gesendet: Montag, 20. Februar 2012 19:28
} An: Mu� Wolfgang
} Betreff: [Microscopy] A protocol for preparation of motor end-plates
} for SEM [pre-synaptic portion!]
}
} ----------------------------------------------------------------------------
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}
} Dear Listers,
}
} I am in a need of a proven protocol for preparing motor end-plates
} (NMJ) for SEM. I've found several that employ use of 8N HCL and then
} collagenase, but I find this treatment to harsh for my gentle muscle.
} Moreover, these papers show only the post-synaptic part, while I need
} to focus on the presynaptic part.
} Any advice would be welcomed.
}
} As always, greetings from Seville,
} Josif
}
} Dr.�Josif�Mircheski
} _______________________________________________________________________________
} Instituto de Biomedicina de Sevilla (IBIS), Lab 108
} Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de
} Sevilla
} Avda. Manuel Siurot S/N�
} 41013 Sevilla
} Spain
}
} Phone:+34-955923045
} e-mail:�jmircheski-at-us.es
} web: www.ibis-sevilla.es
}
}
}
}
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Hi,

A colleague has been attempting to cut thin sections of carbon fibres embedded in Spurr's resin for TEM imaging but with little or no success. Sectioning has been attempted transversely, longitudinally and obliquely all with little or no result. The sections are being cut at room temperature as we don't have a cryo-ultramicrotome.

If anyone has a method for sectioning these fibres at room temperature my colleague would appreciate it very much. If cryo is the only way to go then at least I have a good case to put for the funding!

Cheers and thank you very much,

Colin Veitch
Electron Microscopist
CSIRO�Materials Science and Engineering, Geelong Laboratory
PO Box 21, BELMONT, Vic. 3216. Australia.

colin.veitch-at-csiro.au
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Hi Derrick!

My first thought is: it must be essentially dependent on the�fixation step but�you give no precision about the preparation of the sample so it is hard to tell. I suppose that the protist which surface you want to analyse is not hard so you'll probably have to fix and dehydrate. In this case the critical step is probably the optimization of the fixation for both SEM and PCR (i know optimization is hard to achieve on unique specimen, perhaps you can optimize on similar specimen you can grow in culture).
My second thought is: Maybe you can have a look at protocols for FISH on whole cells, they are able to hybridize on fixed cells, it is good start.
http://biowww.net/detail-337.html
http://fds.oup.com/www.oup.co.uk/pdf/pas/12-6-6.pdf�(hybridization of fixed specimen)

Entering "PCR after fixation" in google, the 5th link looks interesting:
http://genome.cshlp.org/content/1/1/46.full.pdf
You may want to look at other links.

Sorry no experience to share but it does not seems impossible to realize.

Good luck
Stephane

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Email:� Name: Derrick Horne

Organization: UBC BioImaging Facility

Title-Subject: [Filtered] Is DNA isolation from SEM prep possible?

Message: Other than through Cryo-SEM techniques, is it possible to isolate DNA from samples prepared
and imaged for the SEM?

Here is the sample type I am thinking about:

Unique non-culturable cells (protista) isolated from the environment. At times there may be only one
or two cells available for both imaging and PCR.

I would love to hear if anyone has any experience trying this. Both positive and negative
experiences may be helpful.

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Dear all,

I am in front of a JEOL JEM 2000 EX II behaving a funny way.

Each time I do change magnification the whole set of shift parameters (condenser and projector) are recalled with a set hat is totally wrong.
I can correct it each time, but as soon as I change magnification, the microscope is recalling the wrong set of parameters.
There is any instruction to store for a certain high voltage a set of shift parameters for all magnifications?
Same for diffraction!
Thanks in advance for any help.

Marco Arienti
www.electronrescue.com

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Hi Marco,

we have a JEOL JEM 1200EX II (so I'm assuming it has the same computer
system)
There are four magnification ranges (M1 to M4) and the one between 10K
and 250K (M3) is the reference range.
(I suspect that the actual numbers may vary between microscopes with
different objective lenses etc..)

I should think that if the alignments change at every magnification then
you have to call Service.

There are adjustments between ranges for beam and image alignment and a
procedure that involves 'teaching' the microscope various settings, like
astigmatism and making it remember the settings. Before you go delving
into the procedure however you must write down the values from the
computer (including perhaps the Hex values). Type PRTEST (space) M2 for
a listing. Make one change at a time and save the result in case you
need to go back.

For example, use image shifts between ranges and PL align within a
range. To save changes you need to enter "learn" mode by first entering
DADJ (space) 1 - then make a change - finish with DADJ (space) 0. This
is a form of programming and you need to start slowly and save often.

For different voltages we use the user free control (UFC) that is
explained well in the manual.

But if you have wrong values, call Service.

Good luck

Rob Keyse

On 2/22/2012 5:37 AM, marienti-at-tiscali.it wrote:
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} Dear all,
}
} I am in front of a JEOL JEM 2000 EX II behaving a funny way.
}
} Each time I do change magnification the whole set of shift parameters (condenser and projector) are recalled with a set hat is totally wrong.
} I can correct it each time, but as soon as I change magnification, the microscope is recalling the wrong set of parameters.
} There is any instruction to store for a certain high voltage a set of shift parameters for all magnifications?
} Same for diffraction!
} Thanks in advance for any help.
}
} Marco Arienti
} www.electronrescue.com
}
} ==============================Original Headers==============================
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--
Robert Keyse
EM Facility
Whitaker Laboratory
5 East Packer Avenue
Bethlehem
PA 18015
USA

Tel. +1 610 758 4283
Fax +1 610 758 4244

==============================Original Headers==============================
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Dear Marco,

It seems that you need to change battery of the memory card of your TEM.
This card is located behind the right console and removable easily after
swich off the TEM; then you will be able to measure voltage of the
battery. Of course all alignment values need to be adjust again when RAM
's contain is lost.
Good luck !
--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

marienti-at-tiscali.it a �crit :
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} Dear all,
}
} I am in front of a JEOL JEM 2000 EX II behaving a funny way.
}
} Each time I do change magnification the whole set of shift parameters (condenser and projector) are recalled with a set hat is totally wrong.
} I can correct it each time, but as soon as I change magnification, the microscope is recalling the wrong set of parameters.
} There is any instruction to store for a certain high voltage a set of shift parameters for all magnifications?
} Same for diffraction!
} Thanks in advance for any help.
}
} Marco Arienti
} www.electronrescue.com
}
} ==============================Original Headers==============================
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}

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

==============================Original Headers==============================
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Dear List,

I look after a pair of Philips CM series TEMs. One of these has recently
developed a beam instability that I have not encountered before. We are
running a CeB6 cathode (over a year old, probably approaching 1,000 hours
use, has be re-positioned once to raise and re-centre the tip), and the
emission current is completely stable. When the beam is focussed at
medium-to-high magnifications, the intensity of the beam fluctuates-
flickers. If the beam is spread just a little, it disappears. This problem
has come and gone over the past few months.

I have hesitated to call out a service engineer as I hoped that changing
the filament might resolve the issue. Is it possible that the filament
could be the cause? I fear a more expensive issue with the HT supply.

Regards,

Ben Micklem

--
Research Support Manager
MRC Anatomical Neuropharmacology Unit, Mansfield Road,
Oxford, OX1 3TH, United Kingdom. Telephone: 01865 271649
{http://www.mrc.ox.ac.uk/}

==============================Original Headers==============================
10, 29 -- From ben.micklem-at-pharm.ox.ac.uk Wed Feb 22 11:07:49 2012
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Hi Ben,
is your condensor aperture mounted on an isolator for reading out the beam-current with a mini-coax connector on the aperture changer?
If yes, check for correct connections / earth continuity.
Or simply a piece of dirt in the beam path (like a small fiber) which gets charged only when the beam hits it.

Best wishes,
Stefan

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

Am 22.02.12 18:12, schrieb ben.micklem-at-pharm.ox.ac.uk:
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} ----------------------------------------------------------------------------
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} Dear List,
}
} I look after a pair of Philips CM series TEMs. One of these has recently
} developed a beam instability that I have not encountered before. We are
} running a CeB6 cathode (over a year old, probably approaching 1,000 hours
} use, has be re-positioned once to raise and re-centre the tip), and the
} emission current is completely stable. When the beam is focussed at
} medium-to-high magnifications, the intensity of the beam fluctuates-
} flickers. If the beam is spread just a little, it disappears. This problem
} has come and gone over the past few months.
}
} I have hesitated to call out a service engineer as I hoped that changing
} the filament might resolve the issue. Is it possible that the filament
} could be the cause? I fear a more expensive issue with the HT supply.
}
} Regards,
}
} Ben Micklem
}
} --
} Research Support Manager
} MRC Anatomical Neuropharmacology Unit, Mansfield Road,
} Oxford, OX1 3TH, United Kingdom. Telephone: 01865 271649
} {http://www.mrc.ox.ac.uk/}
}
}
}
}
}
} ==============================Original Headers==============================
} 10, 29 -- From ben.micklem-at-pharm.ox.ac.uk Wed Feb 22 11:07:49 2012
} 10, 29 -- Received: from relay8.mail.ox.ac.uk (relay8.mail.ox.ac.uk [129.67.1.171])
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} 10, 29 -- Date: Wed, 22 Feb 2012 17:07:46 +0000
} 10, 29 -- Subject: beam instability- causes?
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} 10, 29 -- Thread-Index: AczxhIBVq+3if5caQzy4Q1lyWQxPEg==
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7, 22 -- From stefan.diller-at-t-online.de Wed Feb 22 11:32:51 2012
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Hi Ben,

An HT instability will mainfest itself in the image as well as the beam
since the energy of the electrons through the lenses is varying. I
would suspect an oxide buildup on the inside of the wehnelt. This is a
known problem with LaB6 and could affect CeB6 as well.

As LaB6 evaporates, it will condense on the inside of the wehnelt. If
you look at the wehnelt when changing a LaB6 cathode, you will see a
purplish deposit due to the LaB6 deposits around the wehnelt aperture.
In the presence of oxygen (also water vapor) you start to form the
oxide. The oxide has ~4 orders of magnitude higher evaporation rate
than the boride. Because the oxide is an insulator, a deposit on the
inside of the wehnelt will affect the bias potential around the hole and
cause flickering in the beam. LaB6 is a semiconductor so the deposits
won't affect the beam stability.

This also means that any oxygen or water vapor in the gun area will
cause a significant decrease in cathode life since you will form an
oxide that evaporates quite quickly. Note that the oxide (at least for
La) starts to form around 1100K, so even turning down the cathode a bit
won't help lifetime significantly in the presence of water/oxygen.

We had someone let the cold trap warm up while operating a LaB6
cathode. Within 15-20 minutes he noted beam instability. After pulling
the wehnelt out, I could clearly see a white (oxide) deposit around the
wehnelt aperture. The cathode was basically gone by that point and had
to be replaced. It was an expensive mistake on his part.

Good luck,
Henk

At 2/22/2012 12:09 PM, ben.micklem-at-pharm.ox.ac.uk wrote:
}
}
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} ----------------------------------------------------------------------------
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} Dear List,
}
} I look after a pair of Philips CM series TEMs. One of these has recently
} developed a beam instability that I have not encountered before. We are
} running a CeB6 cathode (over a year old, probably approaching 1,000 hours
} use, has be re-positioned once to raise and re-centre the tip), and the
} emission current is completely stable. When the beam is focussed at
} medium-to-high magnifications, the intensity of the beam fluctuates-
} flickers. If the beam is spread just a little, it disappears. This problem
} has come and gone over the past few months.
}
} I have hesitated to call out a service engineer as I hoped that changing
} the filament might resolve the issue. Is it possible that the filament
} could be the cause? I fear a more expensive issue with the HT supply.
}
} Regards,
}
} Ben Micklem
}
} --
} Research Support Manager
} MRC Anatomical Neuropharmacology Unit, Mansfield Road,
} Oxford, OX1 3TH, United Kingdom. Telephone: 01865 271649
} {http://www.mrc.ox.ac.uk/}
}
}
}
}
}
} ==============================Original Headers==============================
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} 10, 29 -- Received: from relay8.mail.ox.ac.uk (relay8.mail.ox.ac.uk [129.67.1.171])
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} 10, 29 -- +0000
} 10, 29 -- From: Ben Micklem {ben.micklem-at-pharm.ox.ac.uk}
} 10, 29 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
} 10, 29 -- Date: Wed, 22 Feb 2012 17:07:46 +0000
} 10, 29 -- Subject: beam instability- causes?
} 10, 29 -- Thread-Topic: beam instability- causes?
} 10, 29 -- Thread-Index: AczxhIBVq+3if5caQzy4Q1lyWQxPEg==
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--
Hendrik O. Colijn
www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at
once. Lately it doesn't seem to be working."

==============================Original Headers==============================
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Dear All,
I am looking for the "old" version of the EM10 filmholders (the ca 3mm thick ones, 30 holders per stack).
I can offer to interchange them for "new" ones (60 holders per stack).

Best wishes,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
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Hello everyone,

I am a PhD student in the ultrastructural analysis with serial section
TEM. Recently, I am looking for good software (if free license) for
alignment my images. I used to use imod (for tomography), however it
doesn't support ssTEM, and Fiji plugin Linear alignment SIRT not very good
between the grids (Seems I need some manual rotation). Would anyone have
good recommendation?

Thanks a lot and have a nice day

Sincerely,

--
Hong ZHAN PhD student
Institute of Biology of the Ecole Normale Superieure Paris(IBENS)
Genetic and Neurobiology of C. elegans /INSERM U.1024
46, rue d'Ulm
Paris, 75005

==============================Original Headers==============================
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Hi All

Henk has the correct area here in that earlier comments pointed out that the
instability went away when the condenser lens was moved away from crossover.
At crossover the virtual source is imaged thus any filament fluctuation
would be noticed. Moving away from crossover the visualisation of the source
is removed.

Contamination in the condenser system could cause illumination instability
but varying kV or spot size would change the effect and help with fault
finding. High kV would reduce a condenser problem or moving to a smaller
spot size would have the same effect.

Hope this helps but Henk I think has the solution.

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide
Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: colijn.1-at-osu.edu [mailto:colijn.1-at-osu.edu]
Sent: 22 February 2012 18:13
To: protrain-at-emcourses.com

Hi Ben,

An HT instability will mainfest itself in the image as well as the beam
since the energy of the electrons through the lenses is varying. I
would suspect an oxide buildup on the inside of the wehnelt. This is a
known problem with LaB6 and could affect CeB6 as well.

As LaB6 evaporates, it will condense on the inside of the wehnelt. If
you look at the wehnelt when changing a LaB6 cathode, you will see a
purplish deposit due to the LaB6 deposits around the wehnelt aperture.
In the presence of oxygen (also water vapor) you start to form the
oxide. The oxide has ~4 orders of magnitude higher evaporation rate
than the boride. Because the oxide is an insulator, a deposit on the
inside of the wehnelt will affect the bias potential around the hole and
cause flickering in the beam. LaB6 is a semiconductor so the deposits
won't affect the beam stability.

This also means that any oxygen or water vapor in the gun area will
cause a significant decrease in cathode life since you will form an
oxide that evaporates quite quickly. Note that the oxide (at least for
La) starts to form around 1100K, so even turning down the cathode a bit
won't help lifetime significantly in the presence of water/oxygen.

We had someone let the cold trap warm up while operating a LaB6
cathode. Within 15-20 minutes he noted beam instability. After pulling
the wehnelt out, I could clearly see a white (oxide) deposit around the
wehnelt aperture. The cathode was basically gone by that point and had
to be replaced. It was an expensive mistake on his part.

Good luck,
Henk

At 2/22/2012 12:09 PM, ben.micklem-at-pharm.ox.ac.uk wrote:
}
}
}
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} Dear List,
}
} I look after a pair of Philips CM series TEMs. One of these has recently
} developed a beam instability that I have not encountered before. We are
} running a CeB6 cathode (over a year old, probably approaching 1,000 hours
} use, has be re-positioned once to raise and re-centre the tip), and the
} emission current is completely stable. When the beam is focussed at
} medium-to-high magnifications, the intensity of the beam fluctuates-
} flickers. If the beam is spread just a little, it disappears. This problem
} has come and gone over the past few months.
}
} I have hesitated to call out a service engineer as I hoped that changing
} the filament might resolve the issue. Is it possible that the filament
} could be the cause? I fear a more expensive issue with the HT supply.
}
} Regards,
}
} Ben Micklem
}
} --
} Research Support Manager
} MRC Anatomical Neuropharmacology Unit, Mansfield Road,
} Oxford, OX1 3TH, United Kingdom. Telephone: 01865 271649
} {http://www.mrc.ox.ac.uk/}
}
}
}
}
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--
Hendrik O. Colijn
www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at
once. Lately it doesn't seem to be working."

==============================Original Headers==============================
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Greetings everyone:

A colleague is looking at the possible purchase of an "EVOS xl core" all in one digital inverted microscope, from "Advanced Microscopy Group". I am not aware of this company or instrument, so I am wondering if anyone else has any experience and/or recommendations regarding the instrument. I value the opinions of this group quite a lot....

Thanks in advance

shea

Dr. S. Shea Miller
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
960 Carling Avenue | 960, avenue Carling
Ottawa, ON K1A 0C6
E-mail Address / Adresse courriel shea.miller-at-agr.gc.ca
Telephone | T�l�phone 613-759-1760
Facsimile | T�l�copieur 613-759-1701
Teletypewriter | T�l�imprimeur 613-773-2600
Government of Canada | Gouvernement du Canada

==============================Original Headers==============================
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Hi Shea,
We have two Evos Fl microscopes, one in our core facility and one in a PC3 facility. They are great for when a user wants to check transfection efficiency or just take a quick look at their samples or culture flasks. It is very easy to use and to train people on, fits in a culture hood and requires virtually no maintenance (perfect for PC3). Since it uses LEDs, there are no lamps to change and the filters are hard coated Semrock filters. If you would like more details contact me offlist.(no commercial interest)

Regards,
Justin.

-----Original Message-----
X-from: Shea.Miller-at-AGR.GC.CA [mailto:Shea.Miller-at-AGR.GC.CA]
Sent: Friday, 24 February 2012 6:05 AM
To: Justin Ross

Greetings everyone:

A colleague is looking at the possible purchase of an "EVOS xl core" all in one digital inverted microscope, from "Advanced Microscopy Group". I am not aware of this company or instrument, so I am wondering if anyone else has any experience and/or recommendations regarding the instrument. I value the opinions of this group quite a lot....

Thanks in advance

shea

Dr. S. Shea Miller
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
960 Carling Avenue | 960, avenue Carling
Ottawa, ON K1A 0C6
E-mail Address / Adresse courriel shea.miller-at-agr.gc.ca
Telephone | T�l�phone 613-759-1760
Facsimile | T�l�copieur 613-759-1701
Teletypewriter | T�l�imprimeur 613-773-2600
Government of Canada | Gouvernement du Canada

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Dear Microscopists,
It is almost Friday and I think this question will
amuse at least some of you. However, I am asking in full earnest. Not
a joke. I happen to have a cylinder of CO2 for critical point drying
that has been here for 3 years or so, untouched. I need to start up
critical point drying again and I am just wondering whether there is
any reason to be suspicious of this "old" CO2? Seems like it is just
a simple gas in a tank, so what could happen? Contamination from the
tank liner? CO2 is a pretty decent solvent. I would be delighted for
people to say "You are crazy! Nothing wrong with it!" I have some one
else's samples coming in and I'd hate to ruin them. So I am being
careful.

Thanks for your thoughts,
Tobias
--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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Tobias,

Chances are good that it would be OK; however, if the specimen is
irreplaceable, then I would order another tank. Around here such a
tank is about $14.

If the procedure gives strange results (and you use the old tank), you
will aways wonder if that was the cause.

--

John J. Bozzola, Ph.D., Professor
Gratefully Retired Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

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Dear Tobias,
I am using the same large CO2 bottle since nearly ten years without any negative happening.

Best wishes,
Stefan

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

Am 24.02.12 04:17, schrieb baskin-at-bio.umass.edu:
} ----------------------------------------------------------------------------
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} Dear Microscopists,
} It is almost Friday and I think this question will
} amuse at least some of you. However, I am asking in full earnest. Not
} a joke. I happen to have a cylinder of CO2 for critical point drying
} that has been here for 3 years or so, untouched. I need to start up
} critical point drying again and I am just wondering whether there is
} any reason to be suspicious of this "old" CO2? Seems like it is just
} a simple gas in a tank, so what could happen? Contamination from the
} tank liner? CO2 is a pretty decent solvent. I would be delighted for
} people to say "You are crazy! Nothing wrong with it!" I have some one
} else's samples coming in and I'd hate to ruin them. So I am being
} careful.
}
} Thanks for your thoughts,
} Tobias

==============================Original Headers==============================
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Hi Tobias,

I have a really big bottle of CO2 since 12 years, no problems.

However, if you have really "critical" specimens, like cutured cells or
sensible tissue, use CO2 of highest purity ("water-free"), though it is
extremely expensive
the standard CO2 contains a lot of water and is useless in CP-drying of
biological specimens.
( I can imagine that the water content in standard CO2 could cause
corrosion within the bottle over the years. Over here in Germany bottles
have to be "mirrored" in the inside by removing valve and thorough
inspection).

greetings,
peter heimann

:
}
}
} ----------------------------------------------------------------------------
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} ----------------------------------------------------------------------------
}
} Dear Microscopists,
} It is almost Friday and I think this question will
} amuse at least some of you. However, I am asking in full earnest. Not
} a joke. I happen to have a cylinder of CO2 for critical point drying
} that has been here for 3 years or so, untouched. I need to start up
} critical point drying again and I am just wondering whether there is
} any reason to be suspicious of this "old" CO2? Seems like it is just
} a simple gas in a tank, so what could happen? Contamination from the
} tank liner? CO2 is a pretty decent solvent. I would be delighted for
} people to say "You are crazy! Nothing wrong with it!" I have some one
} else's samples coming in and I'd hate to ruin them. So I am being
} careful.
}
} Thanks for your thoughts,
} Tobias

--
====================================================

Dr. Peter Heimann Raum: W7-107 / Tel.: +49-(0)521-106-5628
Universitaet Bielefeld Fax: +49-(0)521-106-5654
"Zellbiologie / Cell Biology" W7-107
33501 Bielefeld, Germany www.uni-bielefeld.de/biologie/cellbio

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Hello,

we are a Materials Science Institute and plan to install a CCD-camera
on an older Philips CM200. The microscope is equipped with a Gatan
PEELS 666, which this will be kept. There is a TV-rate camera installed
in off axis position, which can be replaced.
Compatible with the PEELS will be a retractable, a side mounted or an
off axis camera. I heard some doubt about the compatibility of a re-
tractable camera with the old version of the PEELS. So to be on the safe
side we would prefer a side mounted camera or an off-axis version.
I checked the microscope and could not find the 35 mm usually used
for side mounted cameras. So for me the off axis seems to be the most
convenient solution. I found not so many options available for the off-
axis port as for the side mount. I found are two options from TVIPS.
They offer a 1k*1k camera for the off axis port on the CM, which will
replace the TV-camera, and the same camera for a so called near axis
positions, which is basically side mounted on the PEELS. I have never
seen CCD in off- and near axis position. Does anyone have experience
with that kind of cameras?
The other option will be to change the part of the column and install
a side mounted camera. They have a wider field of view but can be
limited in high resolution due to the demagnification. The maximum
magnification of the microscope is 970k, which might be sufficient.
Usually this is never used because on film 380k or 470k are sufficient.

Thanks in advance!

Roland Schierholz
Instituto de Ciencia de Materiales de Sevilla Calle Am�rico Vespucio 49
41092 Sevilla, Espa�a

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Tobias,

I haven't gone as long as some of the other folks, but I routinely
have a tank for 2 years or more without problem.
I agree about the problems of water in CO2 tanks, but there is a
cheaper-in-the-long-run solution: put a no-go water filter or
desiccator in the line. This way you don't need to buy the really
expensive CO2 - you do need a siphon tank - but, I do recommend
getting food-grade siphon CO2. These might have water problems
(nonissue with a filter), but they won't have any of the oils that
can contaminate non-food-grade CO2. (Mind, I'd also use an oil filter
and a particle filter.)

Phil

} Dear Microscopists,
} It is almost Friday and I think this question will
} amuse at least some of you. However, I am asking in full earnest. Not
} a joke. I happen to have a cylinder of CO2 for critical point drying
} that has been here for 3 years or so, untouched. I need to start up
} critical point drying again and I am just wondering whether there is
} any reason to be suspicious of this "old" CO2? Seems like it is just
} a simple gas in a tank, so what could happen? Contamination from the
} tank liner? CO2 is a pretty decent solvent. I would be delighted for
} people to say "You are crazy! Nothing wrong with it!" I have some one
} else's samples coming in and I'd hate to ruin them. So I am being
} careful.
}
} Thanks for your thoughts,
} Tobias
} --
} _ ____ __ ____
} / \ / / \ / \ \ Tobias I. Baskin
} / / / / \ \ \ Professor
} /_ / __ /__ \ \ \__ Biology Department
} / / / \ \ \ 611 N. Pleasant St.
} / / / \ \ \ University of
} Massachusetts
} / / ___ / \ \__/ \ ____ Amherst, MA, 01003
} www.bio.umass.edu/biology/baskin
} Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Dear Colleagues

I would like to add my experience on the CO2 as a possible cause of sample
damaging during preparation for SEM. Until recently I had never detected any
problems with the industrial rate CO2. I use for CPD. Maybe water
contamination as Peter points out, is a problem in countries where the air
is humid, but not in Athens (well, at least the nice climate in Greece
remains unspoiled). The last couple of years I found a shop that refills
the cylinder on the spot. They have a big tank and they use a pump to refill
the cylinder. Since that time I detected some little spots occurring
sporadically on my samples and they look like something oily. Because they
are very small {30 microns they were puzzling me, until I came with the idea
that they are oil droplets coming from the refilling pump.

Has anybody else noticed such a problem? Please have a look at
http://www.eikonika.net/v2/photo_list_nikas.php (it's an animal epithelium).

Have a nice weekend!

yorgos

Dr Yorgos Nikas
Athens Innovative Microscopy
Skra 36 Voula 16673 GREECE

Tel/fax +30 210 8957677
mobile +30 6945 107477
eikonika-at-otenet.gr
www.aim.cat
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What can we use to stain cells for at least the first steps in fixation
for TEM? User needs to see if her nematosomes (maybe-or-maybe-not cells)
really do get pelleted into agar before further processing. I think I
remember toluidine blue will do, but not sure...

Aloha,
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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Tina,

0.01% (or so) neutral red, or eosin will do - the eosin will come out
in the alcohol deydration steps.

Phil

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Email: grazul-at-ccmr.cornell.edu Name: john grazul

Organization: Cornell University

Title-Subject: [Filtered] Job available at Cornell

Message: Good luck! please check out the web-site and follow the instructions for the application,
https://cornellu.taleo.net/careersection/10164/jobdetail.ftl, BTW, great equipment and fantastic
students, working with me is the only downside!
Job Description - Research Supp Spec III (16981)
Job Description Research Supp Spec III-16981

Description
The Shared Facilities at the Cornel Center for Materials Research (CCMR) seeks a highly motivated
individual with significant experience in advanced electron microscopy. The individual will have
primary responsibility for the operation of multiple scanning electron microscopy systems including
a new state of the art Field Emission Scanning Electron Microscope (FESEM), user training and
support, and operation, maintenance and management of service contracts. As part of a growing,
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techniques.

Qualifications
Master's degree in science or engineering, or equivalent education and a minimum of 3 years
experience in electron microscopy operation. The individual should have significant practical
experience in the EM in general and FESEM operation in particular, and a sound understanding of the
associated analytic techniques such as EDS and EBSD. Other desirable skills include: experience in
programming and electronics. Background check may be required. No relocation assistance is provided
for this position. Visa sponsorship is not available for this position.
Cornell University, located in Ithaca, New York, is an inclusive, dynamic, and innovative Ivy
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Cornell University is an equal opportunity, affirmative action educator and employer.

Job
-Research Support
Primary Location
-Ithaca
Organization
-Cornell Ctr for Materls Rsrch
Schedule
-Full-time
Job Type
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Overtime Status
-Exempt Contact Name
-Jamie Washburn
Number of Openings
-1

University Job Title
-Research Support Specialist III
Level
-G
Sector*
-Endowed

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Note added:
Originally in your request there was displayed within the Listserver form an incorrect e-mail-Address (good to have found your personal website!)

Dear Shannon,
it seems that you might not have gotten many replies on your request, since your e-mail-Address has been displayed incorrectly (see below)
Since I am interested in the Hiraoka Grid Staining kit too (have bought some in the late 80ies but discontinued their use) I digged a little bit.

Oh, yes, I have seen that EMS doesn't display Hiraoka any more,
and Polysciences (located in WARRINGTON PA, USA as well as in Europe) has "Discontinued the product 05/17/2011" according to their online catalog.
BUT:
AGAR Scientific.com (located in Cambridge UK) cf.
http://www.agarscientific.com/catalogue/action_catalogue.asp?spx=1&sat=2&saa=11&jumpto=12HK
has the Hiraoka Grid staining kit in their catalogue: # G3720 , NLA HIRAOKA STAINING KIT.
As you need the product in USA, I recommend writing to them whether an order could also delivered in the USA. AGAR Scientific is shown up as a BRAND of ELEKTRON Technology plc, their headquarters is located:

Elektron Technology plc
Broers Building
JJ Thomson Avenue
Cambridge CB3 0FA UK
Tel:�+44(0) 1223 371 000

and according to their website http://www.elektron-technology.com/
(and find there also : {Global Network & Locations} = see http://www.elektron-technology.com/about-us/global-network-locations/ there must be an office in or near Los Angeles/CA. USA I guess)

and their offer "And for all other queries, including any regarding this website, please email to: plc-at-elektron-technology.com"
there might be at least a chance to get your Hiraoka kit also in the USA. Apologize if this only is a "nebulous" thinking/mind of a European colleague, but so do I if I need something fabricated in the USA and might not be allowed to order directly there).

Wish you a pleasant weekend and good luck with your re-order....
Best regards

Wolfgang MUSS
SALZBURG-AUSTRA

Von: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Gesendet: Samstag, 25. Februar 2012 15:28
An: Mu� Wolfgang
Betreff: [Microscopy] Hiraoka Grid Staining Kit
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NOTE ADDED:
wrong E-Mail Address: correct one= modla-at-dbi.udel.edu
cf. http://bioimaging.dbi.udel.edu/staff/shannon-modla
Organization: Delaware Biotechnology Institute

Title-Subject: Hiraoka Grid Staining Kit
Message: I've been using the Hiraoka grid staining plate for years to stain grids for TEM.
Ours are starting to get worn, and I was searching for a vendor that still supplies them.
The only vendors I found were in Europe or Australia.

Is anyone aware of any US vendors that still carry them?

Thanks,
Shannon

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Hi Rick,
I'd be interested in seeing the picture. When you said it broke just
straight across, my first thought was an intergranular fracture of the wire,
which I see from time to time, but the melted tungsten has me curious.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Email: richard.ross-at-allisontransmission.com Name: Rick

Organization: Allison Transmission Inc.

Title-Subject: [Filtered] Low Tungsten Filament Life - Did It Melt?

Message: We just installed a new SEM, FEI Quanta 650 with a tungsten source.
The first filament just
blew at 24 1/2 hours operation, quite short of expectations. This filament
had been installed and
set up for auto saturation by the installing service engineer. Upon
disassembly, the bluing of the
base and Wehnelt appear typical. The ceramic is just slightly off-white. The
filament break is in
the side wire and straight across, no thinning; similar to what Steve
Chapman shows in Diagram 3C of
his "The Life and Death of a Tungsten Hairpin Filament". What alarms me,
though, is that the inside
bend of the hairpin appears filled with material; not knowing otherwise, it
appears to me to be
melted tundsten! I've never seent this before and wonder if it is due to my
mistake or as set-up
issue with the instrument. Spot size was only increased to 7 or 8 (10 is
max.) maybe 10 times during
x-ray analysis. I have photographs of the broken filament, but cannot upload
them to a storage
service because of company restrictions, but would gladly forward them to
anyone that contacts me
directly. Any comments or suggestions are welcome. Rick

Login Host: 12.53.132.4
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Hi All

Title; Academic Director - Otago Centre for Electron Microscopy.

A confirmation path / permanent position, commencing at the Senior Lecturer or Associate Professor level, has become available in the Otago School of Medical Sciences. The Academic Director will provide overall academic leadership and expertise in cryo-transmission electron microscopy and electron tomography in the Otago Centre for Electron Microscopy (0.5 FTE). The Director will also contribute to teaching and graduate supervision while maintaining an internationally recognised research portfolio in a specific Department (i.e. Anatomy, Biochemistry, Microbiology and Immunology, Pharmacology and Toxicology, or Physiology) within the School of Medical Sciences (0.5 FTE). This vacancy has arisen because of the arrival of a new 200kV cryo-transmission electron microscope with electron tomography capability at the University of Otago.

Further information about this position can be found at;

http://www.otago.ac.nz/

Search for current vacancies, then view current vacancies. This position is vacancy 1200247.

Information about the Otago School of Medical Sciences can be found at; http://osms.otago.ac.nz/

Information about the Otago Centre for Electron Microscopy can be found at; http://ocem.otago.ac.nz/index.php#3

If you would like a copy of the Information for Candidates emailed to you then let me know.

Thanks

Allan

Allan Mitchell
Microscopy Otago - Electron Microscopy
c/- Department of Anatomy
Otago School of Medical Sciences
P.O. Box 913
Dunedin
New Zealand

Phone (03) 479 5642 or 479 7301
Fax (03) 479 5086 or 479 7254

EM Centre: http://ocem.otago.ac.nz/
Confocal Centre: http://occm.otago.ac.nz/
NZ Microscopy Society: http://microscopynz.co.nz/

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Hi Richard

My first thought was based on the fact that I never trust auto saturation if
filament life is the issue. Auto saturate will take the heat level to the
plateau of the filament saturation graph in my mind this is overheating. All
modern instruments, unless working at very low kV or pushing the instrument
to the limit, do not need the heating level to be taken to the plateau! We
often work at 50,000X plus under these conditions.

It seemed as if the filament melted in a strange way. The square ended
break is usually when the filament has melted to create large balls on the
end of the break and the vibration from disassembling has broken them away.
On the other hand too heavy a molten ball may have fallen into the V of the
filament creating the structure that you see; gravity is a funny thing!

Ok, problem probably solved but you really need to repeat the experiment,
using auto to prove over heating or manual to prove you can do it better.

Your filament break form is a first for me but totally understandable.

Regards

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide
Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

Email: richard.ross-at-allisontransmission.com Name: Rick

Organization: Allison Transmission Inc.

Title-Subject: [Filtered] Low Tungsten Filament Life - Did It Melt?

Message: We just installed a new SEM, FEI Quanta 650 with a tungsten source.
The first filament just
blew at 24 1/2 hours operation, quite short of expectations. This filament
had been installed and
set up for auto saturation by the installing service engineer. Upon
disassembly, the bluing of the
base and Wehnelt appear typical. The ceramic is just slightly off-white. The
filament break is in
the side wire and straight across, no thinning; similar to what Steve
Chapman shows in Diagram 3C of
his "The Life and Death of a Tungsten Hairpin Filament". What alarms me,
though, is that the inside
bend of the hairpin appears filled with material; not knowing otherwise, it
appears to me to be
melted tundsten! I've never seent this before and wonder if it is due to my
mistake or as set-up
issue with the instrument. Spot size was only increased to 7 or 8 (10 is
max.) maybe 10 times during
x-ray analysis. I have photographs of the broken filament, but cannot upload
them to a storage
service because of company restrictions, but would gladly forward them to
anyone that contacts me
directly. Any comments or suggestions are welcome. Rick

Login Host: 12.53.132.4
---------------------------------------------------------------------------

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Email: wbcq-at-wbcq.com {mailto:wbcq-at-wbcq.com} Name: Allan Weiner

Title-Subject: [Filtered] Early RCA type electron microscope

Message: Have been looking for an early RCA type electron microscope to ad to our collection. Anyone
know of one available?

Allan

Login Host: 68.205.165.74

Allan
We have one of these in storage ( it used to be on display in our lobby)
Can you be more specific about
How you intend to use it, if you are willing to pay for it and if you are willing to pay to have it shipped to where ever?

Bob
Dr. Robert J. Schmitz
Associate Professor of Anatomical Sciences
Department of Biology
University of Wisconsin-Stevens Point
800 Reserve Street, 380 TNR Bld
Stevens Point, WI 54481
715-346-2420
Email: rschmitz-at-uwsp.edu
http://www4.uwsp.edu/biology/faculty/RSchmitz/

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Dear List:

The New York Structural Biology Center has an LKB Bromma 2088 Ultratome
V in need of a new home. The machine was serviced last year but hasn't
been used in a while. It has all its attachments and parts and would be
great for spare parts. The prospective new home pays for shipping.
Please reply if you're interested.

Thanks,

Mariena

--
Mariena Silvestry Ramos, PhD
CEM Scientist
New York Structural Biology Center
212-939-0660 ext. 9322
msilvestry-at-nysbc.org

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Hi All,
I am a graduate student from Macquarie University, and I was wondering
if anyone has a pyrite FeS2 specimen (purity/homogeneity checked by WDS
or other means) that I can have/borrow as a SEM EDS standard? I can
unfortunately not afford to buy a pyrite standard as part of a set of
commercially available EDS standards. If anyone know any commercially
available single mineral EDS standard, I would also appreciate knowing.
Thank you for any help you can provide.
Sincerely,
Amy

--
Amy P. Chen
Building E7A, Room 831
Department of Environment& Geography - Environmental Science
Macquarie University
Balaclava Road, North Ryde
New South Wales, 2109
Australia
+61 (02) 9850-8318 (office)
amy.chen-at-mq.edu.au
http://www.envsci.mq.edu.au/index.html

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Hi

Now I have had chance to see a picture of the filament I see a different
situation to what I envisaged.

This filament does not show any thinning at all so the break was not due to
the auto saturation; I believe the filament was under stress. This is more
of a fracture than the heat generated thinning and failure that we normally
see. So relax, the evidence does not point to auto saturation as the guilty
party but more towards an inbuilt stress. A crack in the filament which was
under a twisting stress (note the offset of the break) such that with a
small amount of thinning the stress was sufficient to cause the facture.

Had we seen a strong taper in the opposite sides of the filament and a
bigger space between the two broken ends, then my original diagnosis would
have fitted.

Great fun and very interesting but you need to blame the filament
manufacturer for the filament failure; no errors at your end!

Regards

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide
Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: Richard A. Ross [mailto:richard.ross-at-allisontransmission.com]
Sent: 27 February 2012 14:45
To: protrain-at-emcourses.com

Hi Richard

My first thought was based on the fact that I never trust auto saturation if
filament life is the issue. Auto saturate will take the heat level to the
plateau of the filament saturation graph in my mind this is overheating. All
modern instruments, unless working at very low kV or pushing the instrument
to the limit, do not need the heating level to be taken to the plateau! We
often work at 50,000X plus under these conditions.

It seemed as if the filament melted in a strange way. The square ended
break is usually when the filament has melted to create large balls on the
end of the break and the vibration from disassembling has broken them away.
On the other hand too heavy a molten ball may have fallen into the V of the
filament creating the structure that you see; gravity is a funny thing!

Ok, problem probably solved but you really need to repeat the experiment,
using auto to prove over heating or manual to prove you can do it better.

Your filament break form is a first for me but totally understandable.

Regards

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide Tel +44
(0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

Email: richard.ross-at-allisontransmission.com Name: Rick

Organization: Allison Transmission Inc.

Title-Subject: [Filtered] Low Tungsten Filament Life - Did It Melt?

Message: We just installed a new SEM, FEI Quanta 650 with a tungsten source.
The first filament just
blew at 24 1/2 hours operation, quite short of expectations. This filament
had been installed and
set up for auto saturation by the installing service engineer. Upon
disassembly, the bluing of the
base and Wehnelt appear typical. The ceramic is just slightly off-white. The
filament break is in
the side wire and straight across, no thinning; similar to what Steve
Chapman shows in Diagram 3C of
his "The Life and Death of a Tungsten Hairpin Filament". What alarms me,
though, is that the inside
bend of the hairpin appears filled with material; not knowing otherwise, it
appears to me to be
melted tundsten! I've never seent this before and wonder if it is due to my
mistake or as set-up
issue with the instrument. Spot size was only increased to 7 or 8 (10 is
max.) maybe 10 times during
x-ray analysis. I have photographs of the broken filament, but cannot upload
them to a storage
service because of company restrictions, but would gladly forward them to
anyone that contacts me
directly. Any comments or suggestions are welcome. Rick

Login Host: 12.53.132.4
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Greetings Fellow Microscopists,

The Midwest Microscopy and Microanalysis Society will hold its first meeting of 2012 on Friday, March 23rd, at Northwestern University in Evanston, IL. The topic is quantitative microscopy and image analysis, and we have an excellent line-up of speakers. A preliminary announcement can be found on our website under Meetings:

www.midwestmicroscopy.org

Please join us for an informative program and a chance to meet with your local colleagues and vendors. We look forward to seeing you there!

Elaine Schumacher
M3S Program Coordinator

*********************************************************************
Elaine F.Schumacher
Senior Research Scientist
McCrone Associates, Inc.
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7417 (fax)
E-mail: eschumacher-at-mccrone.com
Web Site: www.mccrone.com

*********************************************************************
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Hi, Elaine

Can you check the link on the M3S site for the program. �It doesn't
seem to be working and we'd love to see who's on the program.

Thanks,
Barbara Foster
Microscopy/Microscopy Education
www.MicroscopyEducation.com
(972)924-5310

On Tue, Feb 28, 2012 at 8:32 AM, {eschumacher-at-mccrone.com} wrote:
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}
} The Midwest Microscopy and Microanalysis Society will hold its first meeting of 2012 on Friday, March 23rd, at Northwestern University in Evanston, IL. �The topic is quantitative microscopy and image analysis, and we have an excellent line-up of speakers. �A preliminary announcement can be found on our website under Meetings:
}
} www.midwestmicroscopy.org
}
}
} Please join us for an informative program and a chance to meet with your local colleagues and vendors. �We look forward to seeing you there!
}
}
} Elaine Schumacher
} M3S Program Coordinator
}
} *********************************************************************
} Elaine F.Schumacher
} Senior Research Scientist
} McCrone Associates, Inc.
} 850 Pasquinelli Drive
} Westmont, IL �60559-5539 USA
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You can buy loose standards from some of the EM suppliers for about $200 each.

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: amy.chen-at-mq.edu.au [mailto:amy.chen-at-mq.edu.au]
} Sent: Monday, February 27, 2012 9:24 PM
} To: Dusevich, Vladimir
} Subject: [Microscopy] Pyrite FeS2 EDS Standard?
}
}
}
}
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} Hi All,
} I am a graduate student from Macquarie University, and I was wondering
} if anyone has a pyrite FeS2 specimen (purity/homogeneity checked by WDS
} or other means) that I can have/borrow as a SEM EDS standard? I can
} unfortunately not afford to buy a pyrite standard as part of a set of
} commercially available EDS standards. If anyone know any commercially
} available single mineral EDS standard, I would also appreciate knowing.
} Thank you for any help you can provide.
} Sincerely,
} Amy
}
} --
} Amy P. Chen
} Building E7A, Room 831
} Department of Environment& Geography - Environmental Science Macquarie
} University Balaclava Road, North Ryde New South Wales, 2109 Australia
} +61 (02) 9850-8318 (office)
} amy.chen-at-mq.edu.au
} http://www.envsci.mq.edu.au/index.html
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We are looking for spare parts for an old Wild M5 Microscope,
particularly a rack and pinion focusing mechanism.

Please contact Dr. Peter Bryant {pjbryant-at-uci.edu} directly if you can help!
Thanks

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Have you checked out the Lehigh courses: http://www.lehigh.edu/microscopy/

Mike Bode, Ph.D.
ResAlta Research Technologies Corp.

2102 Beech Ct
Golden, CO 80401
USA

p: +1-303-234-9270
f: +1-303-202-6350
Mike.Bode-at-ResAltaTech.com
www.ResAltaTech.com

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Email: serene_ng-at-dsi.a-star.edu.sg Name: Serene Ng

Organization: Data Storage Institute

Title-Subject: [Filtered] Course for image analysis

Message: I know there are quite a few TEM hands-on operation courses.
Are there any recommended courses for S/HR/TEM image analysis and
interpretation?

Thank you!

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Lee,

I know the piece of apparatus that you have in mind for I have seen one in the past.
I believe that the old glass blowers that are no longer employed by universities used to make them. I have never seen them in a catalogue but know those men had great skills that are being lost.

At U of P there used to be a glass blower who had magnificent glass works hanging on the walls of his workshop. I saw them in the early '90s when he ground my Denton DV502 bell jar back into perfect shape. That time I was asked to assist him for he no longer had a helper. I believe that he learned the trace from his father whom he had replaced when he retired.

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E - Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}

Opinions and experiences related are those of Pat Connelly and do not represent the NIH. This message is not confidential and can be freely shared and reproduced.

-----------------------------------------------

Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar creates films of verying
thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

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Any glassblower should be able to make the gadget you describe for a
very nominal cost. Check with your University glassblowing shop.
--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

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Hello Lee,

What you described is a buret. Check chemical supply houses for a
large volume buret that could accommodate a glass slide. You should be
able to find one, but it will be expensive.

If you have a glassblower at your institution, he could add a stopcock
at the bottom of a graduated cylinder.

--

John J. Bozzola, Ph.D., Professor
Gratefully Retired Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

On Tue, Feb 28, 2012 at 6:49 PM,
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} Title-Subject: [Filtered] apparatus for formvar-coating slides
}
} Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
} The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
} a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
} gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
} didn't know where it came from.
} I have found that dipping and withdrawing the slides from a staining jar creates films of verying
} thickness.
} I've looked in the EM suppliers' catalogs, to no avail.
} Any ideas?
} thanks much !!!
} Lee
}
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Lee,

Just thought of other possibilities: large glass syringe (also called
CT High Pressure Syringe) used in angiography, or a short stubby
chromatography column.

Lastly, try contacting some EM supply houses, like EMS. They may be
able to have one fabricated for you.

--

John J. Bozzola, Ph.D., Professor
Gratefully Retired Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

On Tue, Feb 28, 2012 at 7:42 PM, {bozzola-at-siu.edu} wrote:
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} Hello Lee,
}
} What you described is a buret. Check chemical supply houses for a
} large volume buret that could accommodate a glass slide. You should be
} able to find one, but it will be expensive.
}
} If you have a glassblower at your institution, he could add a stopcock
} at the bottom of a graduated cylinder.
}
}
} --
}
} John J. Bozzola, Ph.D., Professor
} Gratefully Retired Director of IMAGE
} Integrated Microscopy & Graphics Expertise
} Southern Illinois University
} Carbondale, IL 62901
}
}
} On Tue, Feb 28, 2012 at 6:49 PM,
} {microscopylistserver-noreply-at-microscopy.com} wrote:
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} } Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould
} }
} } Organization: Weill Cornell Medical College
} }
} } Title-Subject: [Filtered] apparatus for formvar-coating slides
} }
} } Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
} } The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
} } a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
} } gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
} } didn't know where it came from.
} } I have found that dipping and withdrawing the slides from a staining jar creates films of verying
} } thickness.
} } I've looked in the EM suppliers' catalogs, to no avail.
} } Any ideas?
} } thanks much !!!
} } Lee
} }
} } Login Host: 157.139.44.204
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} } ==============================Original Headers==============================
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} 9, 27 -- Subject: Re: [Microscopy] viaWWW:apparatus for formvar-coating slides
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9, 24 -- From bozzola-at-siu.edu Tue Feb 28 20:04:47 2012
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Several have given you some good ideas. I have two for you.

First, the quick and inexpensive. In the Kodak Research Labs we have
for many years used a very inexpensive system that in my opinion works
as well as the more complicated devices (we have used a couple of them
as well). A 100 mL graduated cylinder is the perfect diameter for a
1x3 in microscope slide. An alligator clip with a stiff bare wire (Cu
ground wire is great) is easily bent to hold the slide over the liquid
to drain for a prescribed time in a solvent-saturated environment to
dry. During drying, we cover the cylinder with a 100 mL beaker. The
longer the drain time, the thinner and more uniform the film. Once you
tune your conditions, the films are quite reproducible. The graduated
cylinders are easily cleaned. We store the stock solution in brown
glass bottles.

In "the good old days" we had at least 6 microscopists doing TEM and
went through a lot of substrates. We found it most economical to make
our own stock solutions and filter them through a Millipore filter. I
am the last microscopist doing TEM - and also do SEM, image
analysis..., and simply can't justify the time to make substrates - I
buy them.

If you really want a glass apparatus and can't find a commercial one
and don't have access to a glass shop at Cornell (that would surprise
me), we still have a full glass shop in the Research Labs here. The
glassblowers are in the same division as our Analytical Labs. Since we
do contract work, you could contact me directly and I could get our
glassblower to make one for you. It is not my intention to promote our
services but just to offer an option to a colleague with a need.

Best regards,
John

On Tue, Feb 28, 2012 at 7:48 PM,
{microscopylistserver-noreply-at-microscopy.com} wrote:
--------------------------------------------------------------------------
}
} Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould
}
} Organization: Weill Cornell Medical College
}
} Title-Subject: [Filtered] apparatus for formvar-coating slides
}
} Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
} The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
} a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
} gravity drainage of the formvar. �I've seen this in other labs, but when asked, the people there
} didn't know where it came from.
} I have found that dipping and withdrawing the slides from a staining jar creates films of verying
} thickness.
} I've looked in the EM suppliers' catalogs, to no avail.
} Any ideas?
} thanks much !!!
} Lee

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7, 36 -- From jrminter-at-gmail.com Tue Feb 28 21:13:01 2012
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Hi Lee,

We purchased a casting device from Ernest Fuller, but that company seems to have disappeared since I last looked.

One other option you could explore is to set up a peristaltic pump so that when it is turned on it slowly pulls on a string. You then attach the string to the casting slide using a bulldog clamp and immerse the slide into a regular Coplin jar, or beaker, containing the formvar solution. Place the peristaltic pump above the formvar solution and switch it on. You will be able to regulate the rate the glass slide is lifted out of the solution using the peristaltic pump, and get a uniform formvar film.

Paul.

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Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar creates films of verying
thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

Login Host: 157.139.44.204
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Hi all,

I would like to hear from anyone who has experience with the SkyScan
Micro-CT attachment for SEM. This attachment uses an X-rays to generate
images for 3-D reconstruction. We would like to hear about what type of
sample have been tried with this and how successful the imaging was. There
are units available with different resolutions so this will play into the
decision if we decide to purchase.

Thanks,
Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Note: still looking for replacement!!
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

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7, 28 -- From: "Sherman, Debra M" {dsherman-at-purdue.edu}
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A bit more digging shows that this is listed commercially here:

http://www.emsdiasum.com/microscopy/products/grids/accessories.aspx

Part Number 71305-01

It would have been easy to miss....

I have no financial interest in Electron Microscopy Sciences; I have
used many of their products over the years and the quality and support
has been good.

Best regards,
John

} On Tue, Feb 28, 2012 at 7:48 PM,
} {microscopylistserver-noreply-at-microscopy.com} wrote:
} --------------------------------------------------------------------------
} }
} } Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould
} }
} } Organization: Weill Cornell Medical College
} }
} } Title-Subject: [Filtered] apparatus for formvar-coating slides
} }
} } Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
} } The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
} } a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
} } gravity drainage of the formvar. �I've seen this in other labs, but when asked, the people there
} } didn't know where it came from.
} } I have found that dipping and withdrawing the slides from a staining jar creates films of verying
} } thickness.
} } I've looked in the EM suppliers' catalogs, to no avail.
} } Any ideas?
} } thanks much !!!
} } Lee
}
}
} ==============================Original Headers==============================
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Hello all

While we are on the subject of casting formvar films.....I have a query
to do with making thicker or thinner films, or perhaps stronger or
weaker films (which maybe is the same thing?). I have recently started
doing a lot of negative staining, some research some diagnostic, with a
variety of specimens, from purified proteins to suspensions of faeces.
Sometimes I would like a thin film so that I can get good structural
detail, while at other times I need a stronger film that will stand up
to a dirtier sample and/or more manipulations of the grid. I'm happy
with my thin films, but I'm struggling to reproducibly make decent
stronger ones (all are carbon-coated before use).

I understand that changing the speed at which the glass slide is
withdrawn from the formvar solution changes the thickness of the formvar
(slower equals thinner). You can also change the thickness by changing
the concentration of the formvar solution that you are using (which is
the method I am currently using to try to make stronger films). My
question is; do changes in these two factors produce the same kind of
qualitative change? To put it another way, If I want a stronger film am
I better off changing my casting speed or the concentration of plastic
in my casting solution...or will I get essentially the same outcome
(theoretically at least) either way? Also does the solvent used to
dissolve the plastic (eg I have seen both chloroform and
ethylenedichloride used for formvar) play a significant role in the
quality/strength of the film cast?

Alternatively, have people out there got good (or bad) experience of
other plastics that might be more suitable as a stronger substrate for
negative staining work that needs resilience/robustness and decent
imaging but not necessarily ultimate resolution? Please feel free to
reply off the list.

Many thanks

Matthew
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For those who weren't able to access it yesterday, the link to the program for the March meeting is now working. Thanks to those who made me aware of the problem, and I apologize for the inconvenience.

http://www.midwestmicroscopy.org/meetings.htm

Elaine

-----Original Message-----
X-from: Elaine F. Schumacher
Sent: Tuesday, February 28, 2012 8:24 AM
To: 'microscopy-at-microscopy.com'

I believe what you want is carried by Electron Microscopy Sciences.

Film Casting Device An all glass apparatus. It casts uniformly thin films of parlodion,
formvar, or butvar directly onto 1x3 microscope slides. The film casting
solution can be used repeatedly. A built-in fine-pressure-release valve
helps control the speed of drainage. The thickness of the film is
controlled by the concentration of the film solution and the rate of the
drainage. The unit requires 100 mls of film casting solution to start.
Cat. # 71305-01 Complete Film Casting Device

I have no connection to EMS just a happy customer.

Ruth Yamawaki
Stanford University
Department of Comparative Medicine

-----Original Message-----
} From: microscopylistserver-noreply-at-microscopy.com
[mailto:microscopylistserver-noreply-at-microscopy.com] Sent: Tuesday, February 28, 2012 4:57 PM
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Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides
(for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of
about 50-100 ml capacity and a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at
the bottom to allow gravity drainage of the formvar. I've seen this in other labs, but when
asked, the people there didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar
creates films of verying thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

Login Host: 157.139.44.204
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Hi Lee,

Try the URL http://www.cardinal.com/us/en/distributedproducts/ASP/2094-250.asp?cat=laboratory

Cole Parmer: large volume buret with stopcocl [probably redundant].

Hope this helps,

Fred Monson

P.S. I have my own apparatus which does not include a buret. Carrying a long, expensive thing with
a breakable tip around was never apealing, so I have my setup in a reasonable box. Everything is
easily replaced.

________________________________________
} From: microscopylistserver-noreply-at-microscopy.com [microscopylistserver-noreply-at-microscopy.com]
Sent: Tuesday, February 28, 2012 8:00 PM
To: Monson, Frederick

This Question/Comment was submitted to the Microscopy Listserver
using the WWW based Form at http://www.microscopy.com/MLFormMail.html
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Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar creates films of verying
thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

Login Host: 157.139.44.204
---------------------------------------------------------------------------

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Try looking for a Film Casting Device at Electron Microscopy Sciences (EMS Diasum) That is where I
got mine recently.

Good luck,

Lita Duraine
HHMI- Bellen Lab

-----Original Message-----
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To: Duraine, Lita

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Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar creates films of verying
thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

Login Host: 157.139.44.204
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Hi Judy,
We've recently demoed a circular polarization system from Nikon and are about to demo one from
Olympus as well. Kira

Kira Lathrop, MAMS
Research Instructor
University of Pittsburgh
Eye and Ear Institute
203 Lothrop Street Rm 1026
Pittsburgh, PA 15261
lathropkl-at-upmc.edu
412-647-3492

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} While we are on the subject of casting formvar films.....I have a query
} to do with making thicker or thinner films, or perhaps stronger or
} weaker films (which maybe is the same thing?).

...
} imaging but not necessarily ultimate resolution? Please feel free to
} reply off the list.
}
} Many thanks
}
} Matthew
========

Hi Matthew,

I just wonder why are you asking to reply off the list.
I am sure that not just I but some other people also will be interested to see discussion.
The listserver is a wonderful educational tool; I owe it a lot and wish to continue my learning.

Regards,

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

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Greetings,
While we are on the subject of Formvar, I would appreciate
hearing about the differences between Formvar and Butvar. How do they
compare? What motivates using one vs the other??

Thanks!
Tobias
--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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Tobias - Butvar is purported to be more hydrophilic, more adhesive, stronger and more translucent. And presumably able to leap tall buildings in a single bound. Your mileage may vary. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
Sent: Wednesday, February 29, 2012 10:10 AM
To: Phillips, Thomas E.

Greetings,
While we are on the subject of Formvar, I would appreciate hearing about the differences between Formvar and Butvar. How do they compare? What motivates using one vs the other??

Thanks!
Tobias
--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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Hi

Looking for info on what to tell my students regarding current center vs voltage center.

I have often settled on current center because, for me, I would rather have the image stay in the center when focusing. I have been told that the cc and vc seldom are exactly the same and the operator should pick one or the other for alignment.

What is the general practice in labs? Try for both, pick one, or punt?

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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Lee

another alternative is to look for the more standadrd seperating funnel used for separating solvent phases. These are generally enclosed at the top but if you can find one of the correct dimensions with straight sides and can source a friendly glass-blower to remove the top and clean it up you have a ready made film casting device. The funnel is usually easy - the glassblower more difficult.

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: microscopylistserver-noreply-at-microscopy.com [microscopylistserver-noreply-at-microscopy.com]
Sent: 29 February 2012 13:41
To: Malcolm Haswell

Try looking for a Film Casting Device at Electron Microscopy Sciences (EMS Diasum) That is where I
got mine recently.

Good luck,

Lita Duraine
HHMI- Bellen Lab

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Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar creates films of verying
thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

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A former electron microscopist here at UCSF, Toni Milroy, was a big
proponenet of Butvar but I have found that it is more susceptible to
drift in the electron beam than Formvar. I also had some excellent
sample supports made with a solution of ULTEM in chloroform which was
used in the John Sedat lab after extensive testing. Unfortunately, I
have not been able to find the source of that ULTEM and there are many
source of varying quality. None of the ones I tried worked
satisfactorily--usually they were dirty or holey ostensibly from
contaminents. So I feel a bit schizophrenic jumping from one
formulation to another when problems arise. I cannot afford to pay $30
per grid or even $1 per grid so I sometimes have to prepare multiple
castings of films to get a good result.

Larry

On 2/29/2012 8:45 AM, PhillipsT-at-missouri.edu wrote:
}
}
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}
} Tobias - Butvar is purported to be more hydrophilic, more adhesive, stronger and more translucent. And presumably able to leap tall buildings in a single bound. Your mileage may vary. Tom
}
}
} Thomas E. Phillips, Ph.D
} Professor of Biological Sciences
} Director, Molecular Cytology Core
} 2 Tucker Hall
} University of Missouri
} Columbia, MO 65211-7400
} 573-882-4712 (office)
} 573-882-0123 (fax)
} phillipst-at-missouri.edu
}
} http://www.biology.missouri.edu/faculty/phillips.html
} http://www.biotech.missouri.edu/mcc/
}
}
} -----Original Message-----
} X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
} Sent: Wednesday, February 29, 2012 10:10 AM
} To: Phillips, Thomas E.
} Subject: [Microscopy] formvar vs butvar
}
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} Greetings,
} While we are on the subject of Formvar, I would appreciate hearing about the differences between Formvar and Butvar. How do they compare? What motivates using one vs the other??
}
} Thanks!
} Tobias

--
Larry Ackerman, Specialist
Electron Microscopy Lab
Manager, Diabetes& Endocrinology Research Center Microscopy Core
UCSF, Dept. of Anatomy, Rm S657
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu

415-999-4758

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On Feb 29, 2012, at 9:27 AM, jkrupp-at-deltacollege.edu wrote:

} Looking for info on what to tell my students regarding current
} center vs voltage center.
}
} I have often settled on current center because, for me, I would
} rather have the image stay in the center when focusing. I have been
} told that the cc and vc seldom are exactly the same and the operator
} should pick one or the other for alignment.
}
} What is the general practice in labs? Try for both, pick one, or punt?
}
} Thanks
}
} Jon
}
} Jonathan Krupp
} Delta College
} 5151 Pacific Ave.
} Box 212
} Stockton, CA 95207
} 209-954-5284
} jkrupp-at-deltacollege.edu

Hi Jon,
I'd have to go to one of the TEM books to define current and voltage
centers, but it depends on your scope (and sometimes preference) what
to align. On the primitive end, the HVEM current center was aligned
by moving the objective lens upper pole piece, and the voltage center
was not aligned, but both the Polara and the Titan from FEI had
electronic alignments for both centers. I am no longer employed by
FEI, but I was a few years ago.
Yours,
Bill

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Hi Jon,

Both are to align the beam to the objective lens.
Generally for materials science lattice imaging, generally near Scherzer focus, the voltage center is done. The high voltage is varied and the beam is tilted to minimize image movement with high voltage change.
For biological imaging and especially with biological tomograms where defocus values vary over many microns, the objective current center must be done. The beam tilt is aligned to minimize image movement with changes in the objective lens.

The two alignments are slightly different and vary from scope to scope. If specified in the original order they can be pretty close to spot on but that is seldom written into the specs.

All the best,
Roseann

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

On Feb 29, 2012, at 9:25 AM, jkrupp-at-deltacollege.edu wrote:

}
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} Hi
}
} Looking for info on what to tell my students regarding current center vs voltage center.
}
} I have often settled on current center because, for me, I would rather have the image stay in the center when focusing. I have been told that the cc and vc seldom are exactly the same and the operator should pick one or the other for alignment.
}
} What is the general practice in labs? Try for both, pick one, or punt?
}
} Thanks
}
} Jon
}
} Jonathan Krupp
} Delta College
} 5151 Pacific Ave.
} Box 212
} Stockton, CA 95207
} 209-954-5284
} jkrupp-at-deltacollege.edu
}
} Find us on Facebook -at-
} Electron Microscopy at SJ Delta College
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Hi all,

I usually prepare my own formvar films in a very "primitive" way: thoroughly
clean the slide (acetone or ethanol) and let it dry for several minutes;
fill in a 50-ml plastic falcon tube with 50 ml of 0.30% formvar in
chloroform; dip the slide fast to some 2/3 of its length into the falcon;
remove the slide very slowly at constant rate by hand.

It needs practise to learn the exact speed of removing the slide by hand.
There are some problems with my method, I know, but it proved to be the
fastest and most convenient way to cast the film and coat the grids. I know
that hand-drawing of the slide produces unequal thickness of the film, but
so far I haven't noticed any thickness-of-film related artefacts produced by
this method. Before coating the grids, I make sure that the film prepared in
that day is good enough by coating a single grid and checking it in the TEM.

0.2% or even 0.25% films proved to be too thin (disintegrating under the
beam).
As far as the film-casting device, the one that is on sale by the EM
distributors proved to be worth one small grant, so I decided to construct
one of my own, easily replaceable.

As far as butvar vs. Formvar, no experience, but I'd like to test and
compare them.

Best,
Josif

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

-----Original Message-----
X-from: larry.ackerman-at-ucsf.edu [mailto:larry.ackerman-at-ucsf.edu]
Sent: Wednesday, February 29, 2012 6:57 PM
To: jmircheski-at-us.es

A former electron microscopist here at UCSF, Toni Milroy, was a big
proponenet of Butvar but I have found that it is more susceptible to
drift in the electron beam than Formvar. I also had some excellent
sample supports made with a solution of ULTEM in chloroform which was
used in the John Sedat lab after extensive testing. Unfortunately, I
have not been able to find the source of that ULTEM and there are many
source of varying quality. None of the ones I tried worked
satisfactorily--usually they were dirty or holey ostensibly from
contaminents. So I feel a bit schizophrenic jumping from one
formulation to another when problems arise. I cannot afford to pay $30
per grid or even $1 per grid so I sometimes have to prepare multiple
castings of films to get a good result.

Larry

On 2/29/2012 8:45 AM, PhillipsT-at-missouri.edu wrote:
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} Tobias - Butvar is purported to be more hydrophilic, more adhesive,
stronger and more translucent. And presumably able to leap tall buildings in
a single bound. Your mileage may vary. Tom
}
}
} Thomas E. Phillips, Ph.D
} Professor of Biological Sciences
} Director, Molecular Cytology Core
} 2 Tucker Hall
} University of Missouri
} Columbia, MO 65211-7400
} 573-882-4712 (office)
} 573-882-0123 (fax)
} phillipst-at-missouri.edu
}
} http://www.biology.missouri.edu/faculty/phillips.html
} http://www.biotech.missouri.edu/mcc/
}
}
} -----Original Message-----
} X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
} Sent: Wednesday, February 29, 2012 10:10 AM
} To: Phillips, Thomas E.
} Subject: [Microscopy] formvar vs butvar
}
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--
Larry Ackerman, Specialist
Electron Microscopy Lab
Manager, Diabetes& Endocrinology Research Center Microscopy Core
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513 Parnassus Ave., Box 0452
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larry.ackerman-at-ucsf.edu

415-999-4758

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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Hi

With the stability of the high voltage being the most sensitive area of an
instrument it is important to have the centre of the high voltage on the
axis of the instrument; voltage alignment.

The historic comment on current and voltage alignment are that the former is
for convenience the latter for resolution. The two should not be far apart
but when pushing an instrument to its limit it is voltage alignment that
results in the higher quality image. Check it out?

Caution, when dealing with biological specimens which may require a wide
range of objective current settings when working over a wide range of
magnifications, then current alignment is the best route as it keeps the
image on the centre of the field of view.

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide
Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
Sent: 29 February 2012 17:18
To: protrain-at-emcourses.com

Hi

Looking for info on what to tell my students regarding current center vs
voltage center.

I have often settled on current center because, for me, I would rather have
the image stay in the center when focusing. I have been told that the cc and
vc seldom are exactly the same and the operator should pick one or the other
for alignment.

What is the general practice in labs? Try for both, pick one, or punt?

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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15, 34 -- Subject: Current v Voltage centering
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30, 23 -- From protrain-at-emcourses.com Wed Feb 29 12:37:42 2012
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30, 23 -- Subject: RE: [Microscopy] Current v Voltage centering
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

To follow up on Steve's comments. One of our TEMs is a BIOTWIN meaning that
the objective lens configuration is designed to increase contrast for
biological samples. In this case the manufacturer's recommendation is to
center using current alignment although voltage alignment could also be used
if desired.

When you purchase a TEM, you often have a choice of objective lens
configuration and the choice is made based on projected use for that
instrument. Thus an instrument designed for low atomic number, poor contrast
samples such as typical biological ones will be different than that chosen
for nano materials or other materials where resolution is more critical.
--
Debby

Debra Sherman, Chief Scientific Officer
DS imaging LLC
Purdue Technology Center
1281 Win Hentschel Blvd
West Lafayette, IN 47906
E-mail: debby.sherman-at-dsimagingllc.com
Mobile: 765-418-8540
Web: www.dsimagingllc.com

} From: "protrain-at-emcourses.com" {protrain-at-emcourses.com}
} Reply-To: "protrain-at-emcourses.com" {protrain-at-emcourses.com}
} Date: Wed, 29 Feb 2012 13:38:49 -0500
} To: Debby Sherman {dsherman-at-purdue.edu}
} Subject: [Microscopy] RE: Current v Voltage centering
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi
}
} With the stability of the high voltage being the most sensitive area of an
} instrument it is important to have the centre of the high voltage on the
} axis of the instrument; voltage alignment.
}
} The historic comment on current and voltage alignment are that the former is
} for convenience the latter for resolution. The two should not be far apart
} but when pushing an instrument to its limit it is voltage alignment that
} results in the higher quality image. Check it out?
}
} Caution, when dealing with biological specimens which may require a wide
} range of objective current settings when working over a wide range of
} magnifications, then current alignment is the best route as it keeps the
} image on the centre of the field of view.
}
} Steve
}
} Steve Chapman FRMS
} Senior Consultant Protrain
} For consultancy and training in electron microscopy world wide
} Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
} www.emcourses.com
}
}
}
} -----Original Message-----
} X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
} Sent: 29 February 2012 17:18
} To: protrain-at-emcourses.com
} Subject: [Microscopy] Current v Voltage centering
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi
}
} Looking for info on what to tell my students regarding current center vs
} voltage center.
}
} I have often settled on current center because, for me, I would rather have
} the image stay in the center when focusing. I have been told that the cc and
} vc seldom are exactly the same and the operator should pick one or the other
} for alignment.
}
} What is the general practice in labs? Try for both, pick one, or punt?
}
} Thanks
}
} Jon
}
} Jonathan Krupp
} Delta College
} 5151 Pacific Ave.
} Box 212
} Stockton, CA 95207
} 209-954-5284
} jkrupp-at-deltacollege.edu
}
} Find us on Facebook -at-
} Electron Microscopy at SJ Delta College
}
}
}
}
}
}
}
}
} ==============================Original Headers==============================
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8, 31 -- From dsherman-at-purdue.edu Wed Feb 29 13:08:29 2012
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Dear Jon,

We had a PhD student who looked into this years ago (Rudiger Meyer). I
remember reading in his PhD thesis that the voltage center (where the beam
energy is wobbled) is mathematically equivalent to finding the coma-free
axis, whereas the rotation center is not. The two always differ by a few
milliradians.

For low-resolution work we do the "Rotation center", i.e. current center,
but insist on using the coma-free axis/volatge center for any sort of
chemical mapping using EFTEM or high resolution imaging (this can be found
in the "Autofilter Tools" panel on FEI instruments, e.g. Tecnai or Titan).
For EFTEM the correct voltage center is important otherwise large image
shifts occur as the HT is adjusted. Otherwise 3-window maps are prone to
losing the edges of the map.

I hope this helps.

Jon

On Feb 29 2012, jkrupp-at-deltacollege.edu wrote:
} Looking for info on what to tell my students regarding current center vs
} voltage center.
}
} I have often settled on current center because, for me, I would rather
} have the image stay in the center when focusing. I have been told that
} the cc and vc seldom are exactly the same and the operator should pick
} one or the other for alignment.
}
} What is the general practice in labs? Try for both, pick one, or punt?
}
} Thanks
}
} Jon

==============================Original Headers==============================
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Not sure I believe the first statement below. It might be useful to read:

The importance of beam alignment and crystal tilt in high resolution electron microscopy
{http://orproxy.lib.utk.edu:2053/science/article/pii/0304399183900062}
Ultramicroscopy, Volume 11, Issue 4, 1983, Pages 263-281
David J. Smith, W.O. Saxton, M.A. O'Keefe, G.J. Wood, W.M. Stobbs

especially section 4. :-)

We would always start with a good voltage centering on our cold FE Hitachi HF-2000, to prep for a following coma-free alignment for high-resolution imaging. After carefully doing a coma-free tilt adjustment, we always found one tilt axis was very close to the value found by voltage center, whereas the second tilt axis was noticeably off by a small amount (don't recall the numbers...).
Larry

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Dear Jon,

We had a PhD student who looked into this years ago (Rudiger Meyer). I
remember reading in his PhD thesis that the voltage center (where the beam
energy is wobbled) is mathematically equivalent to finding the coma-free
axis, whereas the rotation center is not. The two always differ by a few
milliradians.

For low-resolution work we do the "Rotation center", i.e. current center,
but insist on using the coma-free axis/volatge center for any sort of
chemical mapping using EFTEM or high resolution imaging (this can be found
in the "Autofilter Tools" panel on FEI instruments, e.g. Tecnai or Titan).
For EFTEM the correct voltage center is important otherwise large image
shifts occur as the HT is adjusted. Otherwise 3-window maps are prone to
losing the edges of the map.

I hope this helps.

Jon

On Feb 29 2012, jkrupp-at-deltacollege.edu wrote:
} Looking for info on what to tell my students regarding current center vs
} voltage center.
}
} I have often settled on current center because, for me, I would rather
} have the image stay in the center when focusing. I have been told that
} the cc and vc seldom are exactly the same and the operator should pick
} one or the other for alignment.
}
} What is the general practice in labs? Try for both, pick one, or punt?
}
} Thanks
}
} Jon

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Dr. Lawrence F. Allard, FMSA
Distinguished Research Staff Member
High Temperature Materials Laboratory
Microscopy Group
Materials Science and Technology Division
Oak Ridge National Laboratory
1 Bethel Valley Road
PO Box 2008
Oak Ridge, TN 37831-6064
(note: the last 4 lines are sufficient for mailing or overnight courier service)
865-607-1144 (cell)
865-576-5413 (fax)
allardLFjr-at-ornl.gov

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New YorkMicroscopical
Society
Bernard Friedman Memorial Workshops
Use of the Microscope& Polarized Light Microscopy
April 28, May 4, 11, 18, 25, June 1,8,
2012

A basic course on light microscopy which will cover the
following topics:
Theory of microscopy, Kohler
Illumination
Diffraction Theory, Contrast Methods
Polarized light, Phase Contrast,
Interference
Hoffman contrast,
Rheinberg, Dark-field & oblique Illumination

An
advanced course on polarized light microscopy which will cover the following
topics:
The nature of polarized light
The origin and interpretation of
interference colors
Birefringence and crystal
orientation, The Indicatrix
Compensation and variable compensators
Interference figures
and their interpretation

The workshop will
consist of seven consecutive Saturdays of lectures and hands on labs to cover
the theoretical and practical aspects of microscopy. The course instructors are Jan Hinsch formerly of Leica Microsystems, Inc., Dennis O’Leary of Micro-Optical Methods, Mary McCann of McCann Imaging, John Reffner of John Jay College and N.Y.M.S. Instructor Don O'Leary.

WHEN:            April
28, May 4, 11, 18, 25, June 1,8, 2012. 10AM to 4 PM

WHERE:          One Prospect Village Plaza, Clifton, NJ 07013, accessible by
public transportation. Information on car pools and transportation will be
provided.)

COST:             $695 for NYMS members, $725 for non-members
(includes membership) Lunch and course materials are included. Checks made out
to NYMS.

HOW:               Register
using form below. Limited to the first 12 registrants.
                          Send form to:   Don O'Leary, 10 Sampson Street,
Unit 113, Saddle Brook, NJ 07663

FURTHER
INFORMATION: Call D. O'Leary (201) 519-2176,
E-mail: dkoleary-at-verizon.net

PLEASE MAIL THIS APPLICATION WITH YOUR PAYMENT
-------------------------------------------------------------------------
Registration Form Use of the Microscope & Polarized Light
Microscopy
N.Y.M.S. Member_________________ ($695) Non-Member__________($725), April 28 to June
8
Registration for Use of the Microscope only (4
Sessions)
N.Y.M.S. Member_________________ ($395) Non-Member__________($425), April 28 to May
18
Registration for Polarized Light Microscopy Only (4
Sessions)
N.Y.M.S. Member_________________ ($395) Non-Member__________($425), May 18 to June 8
Name______________________________________________________________________
Address____________________________________________________________________
City___________________________State_________________zip____________
Phone
(W)_______________________(H)___________________________

e-mail address___________________.

Please send your application and payment
directly to:

NYMS Spring
2012 Courses
c/o Mel
Pollinger, Treasurer
18-04 Hillery Street
Fair Lawn, NJ 07410-5207

==============================Original Headers==============================
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Sorry for error, Dates are all Saturdays, April 28, May 5,12, 19, 26, June 2, 9 2012

Don O'Leary�

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Hi, Serene

We also provide customized, on-site courses, if that would be of any
help. Please contact me off-line for details.

Best regards,
Barbara Foster
Microscopy/Microscopy Education
www.MicroscopyEducation.com
(972)924-5310

On Tue, Feb 28, 2012 at 7:03 PM, {mike.bode-at-resaltatech.com} wrote:
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Dear Jon,

On FEI/Philips microscopes there are several different alignment options:
- Current center aka rotation center: with the beam spread, the objective lens current is wobbled, any image shift is minimized by adjusting the beam tilt coils.
Good enough for low resolution work, e.g. tomography.

- Coma free alignment: the beam tilt is wobbled some mili-radians around the center value. This causes beam tilt induced astigmatism and defocus (coma) due to the Cs of the objective lens. The difference in focus and astigmatism between the plus and minus beam tilt has to be made symmetric by adjusting the beam tilt coils. This has to be done for both the x and y direction of the image.
For any high resolution work, coma free alignment is the preferred method. It's a quick-and-dirty alternative for a Zemlin tableau, which is superior and used e.g. in the tuning of image Cs correctors.

Many people iterate rotation center and coma free alignment a few times, this is not necessary: rotation center is the rough alignment to bring things close, coma free then is the final fine-tuning.

- Voltage center (only available for energy filters): the accelerating voltage is wobbled, image shift is minimized by adjusting the beam tilt coils. This alignment however affects the entire beam path in the condenser system, the objective lens and the projector system as all lenses are kept constant but the accelerating voltage is changing. There are options for compensation of beam intensity ( = beam size) and beam shift.
This is only needed for convenience when using an energy filter for certain applications where the acceleration voltage is changed.

Best,
Wim

On Feb 29, 2012, at 6:26 PM, jkrupp-at-deltacollege.edu wrote:

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} Looking for info on what to tell my students regarding current center vs voltage center.
}
} I have often settled on current center because, for me, I would rather have the image stay in the center when focusing. I have been told that the cc and vc seldom are exactly the same and the operator should pick one or the other for alignment.
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} What is the general practice in labs? Try for both, pick one, or punt?
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} Thanks
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} Jon
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} Jonathan Krupp
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Hello,
We are using for a long time similar apparatus that Malcolm described. Our device consists of an Erlenmeyer flask, a glass gadget made by our friendly glassblower and a standard latex bulb with an air release valve. The set can be seen here:

http://www2.biomed.cas.cz/~benada/formvar/index.html

Best regards Oldrich

P.S. I'm sorry for quality of the images. They were taken by a mobile phone.

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

On Wednesday 29 of February 2012 18:43:03 you wrote:
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} another alternative is to look for the more standadrd seperating funnel used for separating solvent phases. These are generally enclosed at the top but if you can find one of the correct dimensions with straight sides and can source a friendly glass-blower to remove the top and clean it up you have a ready made film casting device. The funnel is usually easy - the glassblower more difficult.
}
} Malcolm
}
} Malcolm Haswell
} Imaging Suite
} Faculty of Applied Sciences
} University of Sunderland
} SUNDERLAND
} SR1 3SD
} UK
} email: malcolm.haswell-at-sunderland.ac.uk
}
} ________________________________________
} X-from: microscopylistserver-noreply-at-microscopy.com [microscopylistserver-noreply-at-microscopy.com]
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} Subject: [Microscopy] [Filtered] RE: viaWWW:apparatus for formvarcoating slides
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} Try looking for a Film Casting Device at Electron Microscopy Sciences (EMS Diasum) That is where I
} got mine recently.
}
} Good luck,
}
} Lita Duraine
} HHMI- Bellen Lab
}
}
}
} -----Original Message-----
} } From: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: Tuesday, February 28, 2012 7:04 PM
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} Title-Subject: [Filtered] apparatus for formvar-coating slides
}
} Message: I've been trying to find the apparatus for formvar-coating slides (for coating grids).
} The thing I have in mind is a straight-sided separatory funnel/buret of about 50-100 ml capacity and
} a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at the bottom to allow
} gravity drainage of the formvar. I've seen this in other labs, but when asked, the people there
} didn't know where it came from.
} I have found that dipping and withdrawing the slides from a staining jar creates films of verying
} thickness.
} I've looked in the EM suppliers' catalogs, to no avail.
} Any ideas?
} thanks much !!!
} Lee
}
} Login Host: 157.139.44.204
} ---------------------------------------------------------------------------
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}
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Let me ask a couple of questions about calibrating a TEM, or rather verification of calibration.

I calibrate with a diffraction grating replica at low magnification and follow the procedure outlined by Ernest Fullam: I measure the same particle at= several calibrated magnification and use an average value to calibrate my = higher magnification.
So far so good!

Then I use NIST traceable Nanospheres to verify the calibration.

I'm in discussions with our QC guy. He a nice fellow and we both want to improve the measurement values we collect. Unfortunately, my nanosphere average values aren't falling close enough to the publish value.

I'm sure the scope is set up properly and the sample height is correct.

So, my two questions are,
Do other TEM operators verify their magnifications and what are do you consider an acceptable difference? 1%, 8%, one standard deviations or ??????
And
Since the camera seems to be the connecting bridge between the image and measurement, does anyone know of literature or have experience with camera resolution and its affect on measurements?

Thanks for your help.................

Frank

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10, 24 -- From frank_karl-at-ardl.com Thu Mar 1 12:12:33 2012
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On Mar 1, 2012, at 10:24 AM, frank_karl-at-ardl.com wrote:

} Let me ask a couple of questions about calibrating a TEM, or rather
} verification of calibration.
}
} I calibrate with a diffraction grating replica at low magnification
} and follow the procedure outlined by Ernest Fullam: I measure the
} same particle at= several calibrated magnification and use an
} average value to calibrate my = higher magnification.
} So far so good!
}
} Then I use NIST traceable Nanospheres to verify the calibration.
}
} I'm in discussions with our QC guy. He a nice fellow and we both
} want to improve the measurement values we collect. Unfortunately,
} my nanosphere average values aren't falling close enough to the
} publish value.
}
} I'm sure the scope is set up properly and the sample height is
} correct.
}
} So, my two questions are,
} Do other TEM operators verify their magnifications and what are do
} you consider an acceptable difference? 1%, 8%, one standard
} deviations or ??????
} And
} Since the camera seems to be the connecting bridge between the image
} and measurement, does anyone know of literature or have experience
} with camera resolution and its affect on measurements?
}
} Thanks for your help.................

Dear Frank,
It sounds like the difficulty is that the measured mag is not close
to the nominal mag; is that correct? If you repeat the calibrations
either immediately or after several months, are the values for the
measured mags close? I have always calibrated the scope I was working
with either once or twice a year, and there were only minor variations
between calibrations, although there were often relatively large
differences between the nominal and measured mags--often due to the
difference in distance between the CCD and the film plane.
For even a 1k x 1k CCD, if the grating or nanosphere covers a large
part of the image, its image size will be several hundred pixels, so a
difference of a pixel might not be significant. Of course, one should
measure a sufficient number of lines of a grating so that the measured
distance is as many pixels as possible, both to increase image size
and to average over as many grating spacings as possible. If the
point-spread function of the CCD is too broad, the pixel representing
the edge of a grating line (or any other object) will be difficult to
determine, and this could affect the accuracy of your measurement, but
taking Fourier transforms and locating the peaks will overcome this--
again by averaging.
Yours,
Bill

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Hi Hongwen,

The first guess is that you over-exposed the image. If your data is a
signed 16 bit integer, going above 32K counts looks (to the computer)
like a negative number. If you don't spread your beam as you drop mags
or if there is a change in the lens states, you could easily get your
brightness too high.

This is an easy thing to test. Spread your beam with the C2 lens
(intensity) or increase your spot number (C1 lens) to make the image
less bright on the screen. At some point, the image should appear with
normal contrast

Cheers,
Henk

At 3/1/2012 7:38 PM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Email: hongwen.zhou-at-temple.edu Name: Hongwen Zhou
}
} Organization: Temple University
}
} Title-Subject: [Filtered] Images taken by CCD at low magnification inverted
}
} Message: The images were taken at magnifications of 3,000 and below at 80 kV. The electron-dense
} regions appear bright and the electron-transparent regions appear dark. The CCD gain reference
} images are up to date, although they were not taken at the same magnification. No such problems were
} observed at higher magnifications.
} Has anybody encountered the same problem? Any suggestions on how to correct it?
}
} Thanks!
}
} Hongwen Zhou
}
} ===========================
} MRF-at-CST, Temple University
} 1901 N 13 St, Beury Hall 205& 206
} Philadelphia, PA 19122, USA
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--
Hendrik O. Colijn
www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at
once. Lately it doesn't seem to be working."

==============================Original Headers==============================
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Contents Retrieved from Microscopy Listserver Archives
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Hello Frank,

When I was doing QC for commercial vendors, I would always run a NIST
traceable standard every time I ran an unknown. For example, if we
were measuring nanopsheres sized in the 50-200 nm range, I would use a
100 nm standard. All samples were prepared at the same time (same
grids, etc.) and examined in the TEM under identical operating
conditions.

I tried to complete the work in one day. If several days elapsed
between microscopy sessions, then I would reshoot the standard along
with the unknowns.

A very good, reliable standard is the MAGICAL standard. Expensive,
yes, but it is accurate over a very wide magnification range. It could
be used for general calibration of the TEM. When we used the MAGICAL
standard to calibrate the TEM, then measured the NIST standards, the
accuracy was around 2-4%. Not bad, but it could be better. I believe
some of the variability was due to the thickness of the MAGICAL
standard.

Always, whenever we evaluated commercial specimens, we would use NIST
standards. With the NIST standards, the accuracy was better than 1%.

--

John J. Bozzola, Ph.D., Professor
Gratefully Retired Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

On Thu, Mar 1, 2012 at 7:03 PM, {wtivol-at-sbcglobal.net} wrote:
}
}
}
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} On Mar 1, 2012, at 10:24 AM, frank_karl-at-ardl.com wrote:
}
} } Let me ask a couple of questions about calibrating a TEM, or rather
} } verification of calibration.
} }
} } I calibrate with a diffraction grating replica at low magnification
} } and follow the procedure outlined by Ernest Fullam: I measure the
} } same particle at= several calibrated magnification and use an
} } average value to calibrate my = higher magnification.
} } So far so good!
} }
} } Then I use NIST traceable Nanospheres to verify the calibration.
} }
} } I'm in discussions with our QC guy. He a nice fellow and we both
} } want to improve the measurement values we collect. Unfortunately,
} } my nanosphere average values aren't falling close enough to the
} } publish value.
} }
} } I'm sure the scope is set up properly and the sample height is
} } correct.
} }
} } So, my two questions are,
} } Do other TEM operators verify their magnifications and what are do
} } you consider an acceptable difference? 1%, 8%, one standard
} } deviations or ??????
} } And
} } Since the camera seems to be the connecting bridge between the image
} } and measurement, does anyone know of literature or have experience
} } with camera resolution and its affect on measurements?
} }
} } Thanks for your help.................
}
}
} Dear Frank,
} It sounds like the difficulty is that the measured mag is not close
} to the nominal mag; is that correct? If you repeat the calibrations
} either immediately or after several months, are the values for the
} measured mags close? I have always calibrated the scope I was working
} with either once or twice a year, and there were only minor variations
} between calibrations, although there were often relatively large
} differences between the nominal and measured mags--often due to the
} difference in distance between the CCD and the film plane.
} For even a 1k x 1k CCD, if the grating or nanosphere covers a large
} part of the image, its image size will be several hundred pixels, so a
} difference of a pixel might not be significant. Of course, one should
} measure a sufficient number of lines of a grating so that the measured
} distance is as many pixels as possible, both to increase image size
} and to average over as many grating spacings as possible. If the
} point-spread function of the CCD is too broad, the pixel representing
} the edge of a grating line (or any other object) will be difficult to
} determine, and this could affect the accuracy of your measurement, but
} taking Fourier transforms and locating the peaks will overcome this--
} again by averaging.
} Yours,
} Bill
}
}
}
}
} ==============================Original Headers==============================
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==============================Original Headers==============================
11, 26 -- From bozzola-at-siu.edu Thu Mar 1 22:50:13 2012
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Contents Retrieved from Microscopy Listserver Archives
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Changhui,
I can't help you with the first question, but the answer to the second
question is to buy a USB floppy drive for about $20. For example
http://www.newegg.com/Product/Product.aspx?Item=N82E16821103402
These require no power beyond the USB port and work quite well.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Email: changhuilei-at-yahoo.com Name: changhui lei

Organization: U of Houston

Title-Subject: [Filtered] update the computer

Message: Hi, colleagues,

We are going to update our computer which is connected to a 2010F installed
with GIF 2001 (NOT Fas
TEM). The old computer is Gateway PIII Pro M1000. I am seeking a few advice
from those who had
updated the computer to GIF.

1) which kind of computer is good. Gatan suggests Precision T7500 which
costs more than $1800. We do
not want a very great computer as our GIF is not very new. But we hope the
the computer could be
compatible with board and could take images and data at reasonably good
speed.

2) The installation of software. The old software is on some 3.5" floppy
disks. The new computer
usually does not have disk drive. Here I am wondering how to install the
software. I assume it is
not just as simple as copying the files from old computer.

Any suggestion and feedback are highly appreciated.

Changhui

Login Host: 129.7.49.16
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Hi Rick,
Glad some of the info on the website has been useful. I really should spend
some time and update it because there have been a lot of vendor changes.

Your break looks like an intergranular break, or basically a flaw in the
wire and the offset is due to just a little stress present. Sometimes you
can't even find the break without pushing on the filament.

I'm baffled by the drop in the bend. You may have the answer, but I've
never seen it before (but even after almost 35 years servicing these things,
they still provide interest with new things to see).

Hopefully changing your autobias number will take care of it. I understand
the field engineers' thinking. I just had to try and explain to a customer
that the image I had of my strontium ferrite sample at 150kX was probably
going to look significantly better than his largely carbon catalyst
membranes, because I can cheat and use ideal specimens, whereas he's stuck
with real world stuff.

Ken

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: Richard A. Ross [mailto:richard.ross-at-allisontransmission.com]
Sent: Monday, February 27, 2012 9:29 AM
To: Ken Converse

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Email: richard.ross-at-allisontransmission.com Name: Rick

Organization: Allison Transmission Inc.

Title-Subject: [Filtered] Low Tungsten Filament Life - Did It Melt?

Message: We just installed a new SEM, FEI Quanta 650 with a tungsten source.
The first filament just
blew at 24 1/2 hours operation, quite short of expectations. This filament
had been installed and
set up for auto saturation by the installing service engineer. Upon
disassembly, the bluing of the
base and Wehnelt appear typical. The ceramic is just slightly off-white. The
filament break is in
the side wire and straight across, no thinning; similar to what Steve
Chapman shows in Diagram 3C of
his "The Life and Death of a Tungsten Hairpin Filament". What alarms me,
though, is that the inside
bend of the hairpin appears filled with material; not knowing otherwise, it
appears to me to be
melted tundsten! I've never seent this before and wonder if it is due to my
mistake or as set-up
issue with the instrument. Spot size was only increased to 7 or 8 (10 is
max.) maybe 10 times during
x-ray analysis. I have photographs of the broken filament, but cannot upload
them to a storage
service because of company restrictions, but would gladly forward them to
anyone that contacts me
directly. Any comments or suggestions are welcome. Rick

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Happy Friday listers,

Occasionally when I start our Oxford Inca EDS application (v. 4.02) a
dialog box will pop up with the title bar "Application Error" and the
message "Call to Undefined Dynalink". I can dismiss the dialog with no
apparent ill effects - all the EDS functions that I commonly use seem to
run normally. Closing the program and restarting will often bring up the
program normally with no error. Googling around I find references to
this error in the context of Microsoft Office programs, and a dll that
can't be found or loaded. The Oxford software talks to Office in some
ways in the "Report" module (which I never really use), so if my error
is just Word or something tripping over itself I don't have any real
worries. But as usual, I like to ping the collective and see if others
have had the same experience, especially if this is something less
innocuous than a Microsoft error.

Thanks in advance,

Jim

--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

Son, if you really want something in this
life, you have to work for it. Now quiet!
They're about to announce the lottery numbers.
- Homer Simpson

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Greetings Fellow Microscopists,

The Hooke College of Applied Sciences, located in Westmont, IL, is offering a scanning electron microscopy short course March 26-30, 2012.� In addition to lectures, this course emphasizes hands-on training using state-of-the-art equipment.� For further SEM training details and registration information, please follow the link below:

http://www.hookecollege.com/courses/course.asp?COURSE_ID=42

Best regards-
__________________________________________________
Chris Gorman
Hooke College of Applied Sciences
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7412(fax)
www.hookecollege.com

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I don't know the chemical formulation but I found the following:

Butvar B-98
see butvar.com
Acetate Content, (% Polyvinyl Acetate) 2.5 maximum
Butyral Content, (% Polyvinyl Butyral) 80 (approximate)

Formvar
Polyvinyl Formal

re Steve's question: Hey Larry; do you have a good protocol for
preparing the butvar? I aborted after learning that it needed to be
heated, etc.
I have mostly used the solution purchased from EMS at room temperature.
The lab notes I found from Toni did not mentioning heating--just taking
several days to dissolve. I have never used an elevated temperature for
my substrate solutions--perhaps the source of less than perfect films?

Larry

On 3/1/2012 7:08 AM, Tobias Baskin wrote:
} Larry,
} Thanks. Do you happen to know how Butvar and Formvar differ
} chemically??
}
} Tobias
}
} } ----------------------------------------------------------------------------
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} }
} } A former electron microscopist here at UCSF, Toni Milroy, was a big
} } proponenet of Butvar but I have found that it is more susceptible to
} } drift in the electron beam than Formvar. I also had some excellent
} } sample supports made with a solution of ULTEM in chloroform which was
} } used in the John Sedat lab after extensive testing. Unfortunately, I
} } have not been able to find the source of that ULTEM and there are many
} } source of varying quality. None of the ones I tried worked
} } satisfactorily--usually they were dirty or holey ostensibly from
} } contaminents. So I feel a bit schizophrenic jumping from one
} } formulation to another when problems arise. I cannot afford to pay $30
} } per grid or even $1 per grid so I sometimes have to prepare multiple
} } castings of films to get a good result.
} }
} } Larry
} }
} } On 2/29/2012 8:45 AM, PhillipsT-at-missouri.edu wrote:
} } }
} } }
} } } ----------------------------------------------------------------------------
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} } }
} } } Tobias - Butvar is purported to be more hydrophilic, more adhesive,
} } } stronger and more translucent. And presumably able to leap tall
} } } buildings in a single bound. Your mileage may vary. Tom
} } }
} } }
} } } Thomas E. Phillips, Ph.D
} } } Professor of Biological Sciences
} } } Director, Molecular Cytology Core
} } } 2 Tucker Hall
} } } University of Missouri
} } } Columbia, MO 65211-7400
} } } 573-882-4712 (office)
} } } 573-882-0123 (fax)
} } } phillipst-at-missouri.edu
} } }
} } } http://www.biology.missouri.edu/faculty/phillips.html
} } } http://www.biotech.missouri.edu/mcc/
} } }
} } }
} } } -----Original Message-----
} } } X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
} } } Sent: Wednesday, February 29, 2012 10:10 AM
} } } To: Phillips, Thomas E.
} } } Subject: [Microscopy] formvar vs butvar
} } }
} } }
} } }
} } }
} } } ----------------------------------------------------------------------------
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} } }
} } } Greetings,
} } } While we are on the subject of Formvar, I would appreciate
} } } hearing about the differences between Formvar and Butvar. How do
} } } they compare? What motivates using one vs the other??
} } }
} } } Thanks!
} } } Tobias
} }
} } --
} } Larry Ackerman, Specialist
} } Electron Microscopy Lab
} } Manager, Diabetes& Endocrinology Research Center Microscopy Core
} } UCSF, Dept. of Anatomy, Rm S657
} } 513 Parnassus Ave., Box 0452
} } San Francisco, CA 94143
} }
} } larry.ackerman-at-ucsf.edu
} }
} } 415-999-4758
} }
} }
} }
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} } Headers==============================
}
}
} .
}

--
Larry Ackerman, Specialist
Electron Microscopy Lab
Manager, Diabetes& Endocrinology Research Center Microscopy Core
UCSF, Dept. of Anatomy, Rm S657
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu

415-999-4758

==============================Original Headers==============================
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==============================End of - Headers==============================

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Dear Larry,
(dear all interested listers),

since I too was interested in this butvar vs formvar-"problem"
(I also have an own "hand-made" glass ware solution for coating my grids with formvar)
I have just digged a little bit and can offer the following:

Notice: you have to paste the whole following URL in your www.request form (otherwise you won't get the article (:-)) (It seems I have unlocked an otherwise inaccessible pdf)

http://www.google.at/url?sa=t&rct=j&q=butvar%20chemical%20data&source=web&cd=3&sqi=2&ved=0CEgQFjAC&url=http%3A%2F%2F203.158.253.140%2Fmedia%2Fe-Book%2FEngineer%2FMaintenance%2FCoating%2520Technology%2520handbook%2FDK4036ch60.pdf&ei=Sv1RT6OuCcbChAfLzPSRAw&usg=AFQjCNFY13OpFul_KyZG5D2TstplSRL3oQ

via the CTRL/F (find)function you can search for Butvar and Formvar (there are some 30/20 results for Butvar/Formvar in that chapter)

==} pdf of Chapter 60 from: Coatings Technology Handbook, Third Edition (2006)

(Coatings Technology Handbook, Third Edition ISBN: 1574446495
Arthur A. Tracton "Coatings Technology Handbook, Third Edition"
Taylor & Francis | 936 pages | 2005-07-28 | ISBN:1574446495 | PDF | 30MB Hardback �165.00
see: http://www.crcpress.com/product/isbn/9781574446494;jsessionid=nyqmkGJICWG79TODzmge-w** (CRC = Taylor&Francis)

Completely revised and updated, the Coatings Technology Handbook, Third Edition supplies a broad cross-index of the different aspects involved in the discipline. Containing 14 new chapters, the book covers the composition of both organic and inorganic resins, pigments or fillers, and additives, from polymeric fluorocarbons to water borne, solvent-borne, and one hundred percent non-volatile compounds. It examines the testing of raw materials and products and shows dyes used in inks with formulation data. This edition includes a new chapter on specialty pigments for high temperature unique to this book, a chapter on statistical experimentation, a chapter on regulations, and a chapter on formulations with a spreadsheet of formulation calculations. This resource expands your awareness and knowledge of coatings, inks, and adhesives, aids you in problem solving, and increases your level of familiarity with the technology

The URL: { http://www.google.at/#hl=de&gs_nf=1&cp=63&gs_id=2&xhr=t&q=Arthur+A.+Tracton+%22Coatings+Technology+Handbook%2C+Third+Edition%22&pf=p&output=search&sclient=psy-ab&pbx=1&oq=Arthur+A.+Tracton+%22Coatings+Technology+Handbook,+Third+Edition%22&aq=f&aqi=&aql=&gs_sm=&gs_upl=&gs_l=&bav=on.2,or.r_gc.r_pw.r_qf.,cf.osb&fp=e6c8f3eb182c33d8&biw=1229&bih=784 }

finds a pdf displaying the CONTENTS-list.

and:
{ www.qmc.ufsc.br/~minatti/docs/20061/polymer_data_handbook.pdf } : download free
EDITED BY JAMES E. MARK, UNIVERSITY OF CINCINNATI

CTRL/F: (find) butvar: Poly(vinyl butyral) P. R. SUNDARARAJAN pp 910-924, incl. 59 references

If you should have unexpected problems in downloading or retrieving those two articles: I have them in my files....

Best wishes and regards,
and hopefully you have/had a beautiful weekend,

Wolfgang Muss
Salzburg-Austria

} -----Urspr�ngliche Nachricht-----
} Von: larry.ackerman-at-ucsf.edu [mailto:larry.ackerman-at-ucsf.edu]
} Gesendet: Freitag, 02. M�rz 2012 23:53
} An: Mu� Wolfgang
} Betreff: [Microscopy] Re: formvar vs butvar
}
} ---------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------------
} I don't know the chemical formulation but I found the following:

Butvar B-98
} see butvar.com
} Acetate Content, (% Polyvinyl Acetate) 2.5 maximum
} Butyral Content, (% Polyvinyl Butyral) 80 (approximate)

} Formvar
} Polyvinyl Formal

} re Steve's question: Hey Larry; do you have a good protocol for preparing the butvar?
} I aborted after learning that it needed to be heated, etc.
}
} I have mostly used the solution purchased from EMS at room temperature.
}
} The lab notes I found from Toni did not mentioning heating--just taking
} several days to dissolve. I have never used an elevated temperature for
} my substrate solutions--perhaps the source of less than perfect films?
}
} Larry
}
}
} --
} Larry Ackerman, Specialist
} Electron Microscopy Lab
} Manager, Diabetes& Endocrinology Research Center Microscopy Core
} UCSF, Dept. of Anatomy, Rm S657
} 513 Parnassus Ave., Box 0452
} San Francisco, CA 94143
}
} larry.ackerman-at-ucsf.edu
}
} 415-999-4758
}
}
}
} On 3/1/2012 7:08 AM, Tobias Baskin wrote:
} } Larry,
} Thanks. Do you happen to know how Butvar and Formvar differ chemically??
} }
} } Tobias
}
} } } ----------------------------------------------------------------------------
} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} } } ----------------------------------------------------------------------------
}
} } } A former electron microscopist here at UCSF, Toni Milroy, was a big
} proponenet of Butvar but I have found that it is more susceptible to
} drift in the electron beam than Formvar. I also had some excellent
} sample supports made with a solution of ULTEM in chloroform which was
} used in the John Sedat lab after extensive testing. Unfortunately, I
} have not been able to find the source of that ULTEM and there are many
} source of varying quality. None of the ones I tried worked
} satisfactorily--usually they were dirty or holey ostensibly from
} contaminents. So I feel a bit schizophrenic jumping from one
} formulation to another when problems arise. I cannot afford to pay $30
} per grid or even $1 per grid so I sometimes have to prepare multiple
} castings of films to get a good result.
}
} } } Larry
}
}
}
} } } On 2/29/2012 8:45 AM, PhillipsT-at-missouri.edu wrote:
} } } } ------------------------------------------------------------------
} ----------
} } } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
} } } } To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
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} http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } } } ------------------------------------------------------------------
} ----------
}
} Tobias - Butvar is purported to be more hydrophilic, more adhesive,
} stronger and more translucent.
} And presumably able to leap tall buildings in a single bound.
} Your mileage may vary. Tom
}
} } } } Thomas E. Phillips, Ph.D
} } } } Professor of Biological Sciences
} } } } Director, Molecular Cytology Core
} } } } 2 Tucker Hall
} } } } University of Missouri
} } } } Columbia, MO 65211-7400
} } } } 573-882-4712 (office)
} } } } 573-882-0123 (fax)
} } } } phillipst-at-missouri.edu
} } } }
} } } } http://www.biology.missouri.edu/faculty/phillips.html
} } } } http://www.biotech.missouri.edu/mcc/
} } } }
} } } }
} } } } -----Original Message-----
} } } } X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
} } } } Sent: Wednesday, February 29, 2012 10:10 AM
} } } } To: Phillips, Thomas E.
} } } } Subject: [Microscopy] formvar vs butvar
} } }
} } } } ------------------------------------------------------------------
} ----------
} } } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
} } } } To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
} } } } On-Line Help
} http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } } } ------------------------------------------------------------------
} ----------
} } } } Greetings,
} } } } While we are on the subject of Formvar, I would appreciate
} } } } hearing about the differences between Formvar and Butvar. How
} do
} } } } they compare? What motivates using one vs the other??
} } } }
} } } } Thanks!
} } } } Tobias
} } }
} } } --
} } } Larry Ackerman, Specialist
} } } Electron Microscopy Lab
} } } Manager, Diabetes& Endocrinology Research Center Microscopy Core
} } } UCSF, Dept. of Anatomy, Rm S657
} } } 513 Parnassus Ave., Box 0452
} } } San Francisco, CA 94143
} } }
} } } larry.ackerman-at-ucsf.edu
} } }
} } } 415-999-4758
} } }
} } }
} } }
} } } ==============================Original
} Headers==============================
} } } 8, 73 -- From Larry.Ackerman-at-ucsf.edu Wed Feb 29 11:50:05 2012
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} } } 8, 73 -- Date: Wed, 29 Feb 2012 09:49:46 -0800
} } } 8, 73 -- From: "Larry Ackerman" {larry.ackerman-at-ucsf.edu}
} } } 8, 73 -- Reply-to: larry.ackerman-at-ucsf.edu
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==============================Original Headers==============================
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Hi All,

Does anyone out there have an operating procedure for an AMRAY 3200 SEM with
low vacuum capabilities? Would appreciate your sharing off-line if you do
have one.

Thanks,
Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

==============================Original Headers==============================
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Email: reganhll-at-gmail.com Name: Regan

Organization: FMLS

Title-Subject: [Filtered] 300KV Vs 100KV

Message: Dear all,

I was wondering what is the reason for the low contrast of images in a 300KV Microscope compared to
an 100KV microscope. Technically the only difference is that you use a beam of 0.00370nm for 100KV
and 0.00197nm for 300KV.
Theoretically if wavelength changes the resolution increases However its unclear to me how it
affects contrast. there is a small indication however that this affects mean free path by changing
the probability of scattering (p) defined as thickness /wavelength. or something like that.

I would be happy to some one explaining this elaborately. Unfortunately I am biologist not a
physicist, But would be glad to see some equations if well explained. or am I asking too much.

Is it possible to have your 300 kv gun in F30 FEI microscope run in 2 modes 300kv and 100kv?? Does
anybody have a microscope configured and using it in this way.
Greets, J

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Pioloform is much stronger in the beam than Formvar or Butvar. It is soluble in dichloroethane or
chloroform. You should add molecular sieves to trap any water molecules to give you films with less
hole defects. You can prepare films as you do now with formvar or butvar.
Formvar and butvar, so last century.

Rick

-----Original Message-----
} From: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Wednesday, February 29, 2012 8:37 AM
To: Fetter, Richard

I believe what you want is carried by Electron Microscopy Sciences.

Film Casting Device An all glass apparatus. It casts uniformly thin films of parlodion,
formvar, or butvar directly onto 1x3 microscope slides. The film casting
solution can be used repeatedly. A built-in fine-pressure-release valve
helps control the speed of drainage. The thickness of the film is
controlled by the concentration of the film solution and the rate of the
drainage. The unit requires 100 mls of film casting solution to start.
Cat. # 71305-01 Complete Film Casting Device

I have no connection to EMS just a happy customer.

Ruth Yamawaki
Stanford University
Department of Comparative Medicine

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Email: lcgould-at-med.cornell.edu Name: Lee Cohen-Gould

Organization: Weill Cornell Medical College

Title-Subject: [Filtered] apparatus for formvar-coating slides

Message: I've been trying to find the apparatus for formvar-coating slides
(for coating grids).
The thing I have in mind is a straight-sided separatory funnel/buret of
about 50-100 ml capacity and a diameter wide enough to fit a 1 x 3 microscope slide and a stop-cock at
the bottom to allow gravity drainage of the formvar. I've seen this in other labs, but when
asked, the people there didn't know where it came from.
I have found that dipping and withdrawing the slides from a staining jar
creates films of verying thickness.
I've looked in the EM suppliers' catalogs, to no avail.
Any ideas?
thanks much !!!
Lee

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Hi Hongwen,

The first guess is that you over-exposed the image. If your data is a signed 16 bit integer, going
above 32K counts looks (to the computer) like a negative number. If you don't spread your beam as
you drop mags or if there is a change in the lens states, you could easily get your brightness too high.

This is an easy thing to test. Spread your beam with the C2 lens (intensity) or increase your spot
number (C1 lens) to make the image less bright on the screen. At some point, the image should
appear with normal contrast

Cheers,
Henk

At 3/1/2012 7:38 PM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Email: hongwen.zhou-at-temple.edu Name: Hongwen Zhou
}
} Organization: Temple University
}
} Title-Subject: [Filtered] Images taken by CCD at low magnification inverted
}
} Message: The images were taken at magnifications of 3,000 and below at 80 kV. The electron-dense
} regions appear bright and the electron-transparent regions appear dark. The CCD gain reference
} images are up to date, although they were not taken at the same magnification. No such problems were
} observed at higher magnifications.
} Has anybody encountered the same problem? Any suggestions on how to correct it?
}
} Thanks!
}
} Hongwen Zhou
}
} ===========================
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't
seem to be working."

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Dear Regan,

There are two aspects to your question which might be worth separating:

First, is there a fundamental physical difference between a 100kV electron
passing through an atom relative to a 300kV electron?

Second, does the electron optics of the microscope make a difference to the
contrast at these two voltages?

To the first question, the answer is yes, there is a two-fold difference.
The electromagnetic potential of an atom increases the energy of an
electron passing through the atom so that it moves slightly quicker than
electron wave passing out in the vacuum. When it emerges from the atomic
potential (slowing back to its original velocity if the interaction is
elastic) the wave is phase shifted. This phase shift is bigger for a low
beam energy electron (100keV) than it is for a high energy electron
(300keV). The stronger phase shift, the bigger the contrast attainable if
phase contrast imaging modes are used.

Further, the chances of the beam electron scattering inelastically also
rises as the beam energy drops. Specifically, the ionisation cross section
(probability of ionising an atom) rises as the beam energy drops, so that
contrast from inelastically scattered electrons rises too. Therefore, an
'average' sample, looks darker, with more contrast at 100kV than it does at
300kV.

Ray Egerton's book "Electron Energy Loss Spectroscopy in the Electron
Microscope" examines the difference between elastic and inelastic in
chapter 3 (I think). I've found this book extremely useful in the past and
Ray's writing style is clear and accessible (I have no financial interest
in recommending his book!).

The answer to the second question (electron-optical differences) is
probably more subtle. Lower energy electrons are more easily deflected, so
that 100 keV electrons are travelling at slightly higher angles relative to
the optic axis. Aberrations in the lenses (spherical aberration and
chromatic aberration) become more significant, resulting in, for example,
lower point resolution (Cs lambda^3)^(0.25) and larger chromatic blurring.
The lower energy electrons striking your detector (negative/CCD camera)
will have a smaller interaction volume and will create fewer excitations. A
fresh gain reference will be needed at the lower energy plus a fresh set of
magnification calibrations.

In answer to your question about configuring the F30 for 100kV, the answer
is yes you can. The Schottky gun emits electrons with a kinetic energy of
about 4-5 keV into the HT accelerator which can be brought down to as low
as 20 kV. For example, we have a FEI F20 that was aligned for 200 and 100
kV in the factory, but we have since aligned at 80 kV and 40 kV without
difficulty. The only issue we have is the Gatan Imaging Filter (GIF) which
we use to image our TEM images- the GIF usually requires a Gatan engineer
to align it at a set energy ($$!). Fortunately, we were using it in STEM
mode at 80 & 40kV and detecting the electrons wasn't a problem with the
BF/ADF/HAADF detectors.

Yours, Jon

} Title-Subject: [Filtered] 300KV Vs 100KV
}
} Message: Dear all,
}
} I was wondering what is the reason for the low contrast of images in a
} 300KV Microscope compared to an 100KV microscope. Technically the only
} difference is that you use a beam of 0.00370nm for 100KV and 0.00197nm
} for 300KV. Theoretically if wavelength changes the resolution increases
} However its unclear to me how it affects contrast. there is a small
} indication however that this affects mean free path by changing the
} probability of scattering (p) defined as thickness /wavelength. or
} something like that.
}
} I would be happy to some one explaining this elaborately. Unfortunately I
} am biologist not a physicist, But would be glad to see some equations if
} well explained. or am I asking too much.
}
} Is it possible to have your 300 kv gun in F30 FEI microscope run in 2
} modes 300kv and 100kv?? Does anybody have a microscope configured and
} using it in this way. Greets, J

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Rick,
Which Pioloform do you mean? Formvar and Butvar are trade
names for polyvinyl formal and polyvinyl butral. I believe that
Pioloform is a competing trade name and comes with various letters.
For example, Pioloform B is I think the same chemical as Butvar. I am
pretty sure you can get the Pioloform version of Formvar too. So
which Pioloform are you recommending here?

Thanks,
Tobias

}
}
} Pioloform is much stronger in the beam than Formvar or Butvar. It
} is soluble in dichloroethane or
} chloroform. You should add molecular sieves to trap any water
} molecules to give you films with less
} hole defects. You can prepare films as you do now with formvar or butvar.
} Formvar and butvar, so last century.
}
} Rick
}
} -----Original Message-----
} } From: microscopylistserver-noreply-at-microscopy.com
} } [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: Wednesday, February 29, 2012 8:37 AM
} To: Fetter, Richard
} Subject: [Microscopy] [Filtered] RE: viaWWW:apparatus for
} formvarcoating slides
}
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Dear colleagues,

We do work with different kind of users that want to do TEM. Some of them =
want to analyze magnetic material. Magnetic particles and bulk. Last week w=
e had an accident with a sample. The sample was some king of iron alloy pre=
pared by dimpler polishing. A large chunk of if broke inside the OL of our =
TEM-FEG. Very fortunately that piece come out by itself. No need to call th=
e service engineer to disassemble the OL and remove the broken sample.

I was wondering how other EM labs deal with this kind of material. Do you =
have some king of special protocol or just don't accept this kind of sample=
inside the TEM?

We have a service contract for the microscopes, but I did heard from the s=
ervice engineer that using a magnetic material in a TEM is a misuse. And it=
is not covered by our service contract. Some of my colleagues argues that =
we can load any kind of sample in the TEM, and this should be covered by th=
e contract. The fine line in the contract says it is a misuse as the engin=
eer said. Any comment?

What about magnetic particles? We have several users that study nanopartic=
les. Magnetic or doped by magnetic elements like Co. Do you think these pa=
rticle can fly off from the grid and get on the OL? These contamination on =
the OL, can it affect the performance of the TEM?

Thanks,

Carlos

__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

==============================Original Headers==============================
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} Email: reganhll-at-gmail.com Name: Regan
}
} Organization: FMLS
}
} Title-Subject: [Filtered] 300KV Vs 100KV
}
} Message: Dear all,
}
} I was wondering what is the reason for the low contrast of images in
} a 300KV Microscope compared to
} an 100KV microscope. Technically the only difference is that you use
} a beam of 0.00370nm for 100KV
} and 0.00197nm for 300KV.
} Theoretically if wavelength changes the resolution increases However
} its unclear to me how it
} affects contrast. there is a small indication however that this
} affects mean free path by changing
} the probability of scattering (p) defined as thickness /wavelength.
} or something like that.
}
} I would be happy to some one explaining this elaborately.
} Unfortunately I am biologist not a
} physicist, But would be glad to see some equations if well
} explained. or am I asking too much.
}
} Is it possible to have your 300 kv gun in F30 FEI microscope run in
} 2 modes 300kv and 100kv?? Does
} anybody have a microscope configured and using it in this way.
} Greets, J

Dear Regan,
There are two types of contrast that are of concern to biologists, so
I'll restrict my discussion to those. For stained, conventional
plastic sections, the mechanism of contrast is that some electrons are
scattered to large angles primarily by the high-Z atoms of the stain.
These electrons are intercepted by the objective aperture (if no
aperture is inserted, some electrons hit the objective lens lower pole
piece, some are back-scattered, etc.), and these electrons are,
therefore, removed from the image, creating dark regions. For a
sufficiently thin specimen, fewer than half the electrons are
scattered--check the pixel values of the dark vs. light regions of the
image on a CCD camera--so the difference in contrast between 100 kV
and 300 kV is due to the difference in scattering cross-section, with
100 kV electrons being scattered about twice as often as 300 kV
electrons. For thicker or more heavily stained sections, this is not
as much a factor, since even at 300 kV enough electrons are scattered
to give contrast, and scattering from the unstained parts of the
section at 100 kV decreases contrast by increasing the background.
The second type of contrast occurs for unstained specimens such as
frozen-hydrated ones. This is phase contrast, which is caused by the
difference in phase change undergone by electrons traveling through a
slab of ice compared to those that have traveled both through ice and
biological material. The phase change is caused by the electrostatic
potential within the specimen. This will decrease the wavelength of
an electron traveling through a positive potential and increase the
wavelength of an electron traveling through a negative potential. The
relevant equations are E = T + V, or total energy (which is conserved)
is equal to kinetic energy plus potential energy, V = qP, or potential
energy equals the charge times the electrostatic potential, and v/
lambda = ph, or the speed divided by the wavelength, which equals the
circular frequency, is equal to the momentum divided by Plank's
constant. (I think I got my factors of 2pi correct, but someone else
from the list can correct me if I did not.). The changes in
wavelength mean that the phase of each electron emerging from the
specimen will depend on the potential it experiences along its path
through the specimen. These differences are larger compared to 100 kV
than to 300 kV, so for thin specimens, also referred to as weak phase
objects, the contrast will be higher at 100 kV, all other things being
equal. The principle other thing is the coherence of the beam. This
has more of an effect on phase contrast than does accelerating
voltage, and FEG sources are more coherent than LaB6 sources operated
in "tip mode", which are, in turn, more coherent than LaB6 sources
operated in the normal way with emission of electrons from both the
tip and the pyramidal faces of the filament, or, least coherent,
tungsten filaments.
Imperfections in lenses, signal-to-noise ratio in biological images,
and other factors such as the difference between the stain atoms and
the biological features of interest, all have more influence on
resolution than the theoretical maximum achievable due to wavelength.
I hope this gives you a good introduction to contrast. There are any
number of EM books that have several chapters each on contrast
mechanisms and image formation. I suggest you find the one that most
closely matches your knowledge of physics--which should not be
completely lacking in a biologist, especially one who wants to see the
equations.
Finally, most scopes can be operated at several values of
accelerating voltage. All that is necessary is to have good alignment
files stored at the voltages of interest. (older, non-computerized
scopes require realignment every time the HT is changed, but I assume
you do not have one of those.) Every 300 kV instrument I have worked
with has at least factory and installation alignments at 100 kV, so
one can choose the operating voltage pretty freely.
Yours,
Bill

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Dear colleagues,
�
I would need a MSDS for our Agar 100 Resin kit (Epon equivalent).
Perchance someone in the list can help me, Agar scientific did not reply my email.
Thanks in advance

Stephane

==============================Original Headers==============================
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Hi

It's tough working with magnetic samples these days as the instruments do
not have systems that help the operator.

With the development of magnetic materials in the late 1960s the TEM
exchange system allowed the specimen to be lowered into the objective lens
from above, this enabled workers to manipulate the objective focal length,
hence lower the level of magnetic field. This was very easy to perform and
quite good magnetic domain work was achieved. Some manufacturers even had
attachments for this work known as Lorenz (? Spelling) accessories.

Today I suggest using the Z prime or eucentric setting to lift the specimen
as far away from the lower pole piece as possible (focus will move
anticlockwise) to lower the objective lens strength. Another area of gain
is to combine this with a lower accelerating voltage. Be aware the
magnification will be lower than indicated due to the lower objective lens
current.

Try it and see the benefits?

Steve

Steve Chapman FRMS
Senior Consultant Protrain
For consultancy and training in electron microscopy world wide
Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: carlos.inoki-at-lnls.br [mailto:carlos.inoki-at-lnls.br]
Sent: 05 March 2012 02:32
To: protrain-at-emcourses.com

Dear colleagues,

We do work with different kind of users that want to do TEM. Some of them =
want to analyze magnetic material. Magnetic particles and bulk. Last week w=
e had an accident with a sample. The sample was some king of iron alloy pre=
pared by dimpler polishing. A large chunk of if broke inside the OL of our =
TEM-FEG. Very fortunately that piece come out by itself. No need to call th=
e service engineer to disassemble the OL and remove the broken sample.

I was wondering how other EM labs deal with this kind of material. Do you =
have some king of special protocol or just don't accept this kind of sample=
inside the TEM?

We have a service contract for the microscopes, but I did heard from the s=
ervice engineer that using a magnetic material in a TEM is a misuse. And it=
is not covered by our service contract. Some of my colleagues argues that =
we can load any kind of sample in the TEM, and this should be covered by th=
e contract. The fine line in the contract says it is a misuse as the engin=
eer said. Any comment?

What about magnetic particles? We have several users that study nanopartic=
les. Magnetic or doped by magnetic elements like Co. Do you think these pa=
rticle can fly off from the grid and get on the OL? These contamination on =
the OL, can it affect the performance of the TEM?

Thanks,

Carlos

__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

==============================Original Headers==============================
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} realname - Sofia Rydell
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} LOCATION - Copenhagen, Denmark
} SUBJECT_OF_QUESTION - Handheld microscope
} QUESTION - Hello,
} We are considering in investing in a handheld USB microscope at our
} museum. Primary, it will be the archaeologists who will use it, the
} decided magnification is x400.
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} Museum of Copenhagen
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Hello Hongwen,
Can you see the image on TEM main screen at low mags (the same as used for CCD camera recording) in proper contrast?
If not, I would suspect a misaligned LM mode of your TEM. In this case you could see a dark-field image (reverse contrast) of your sample.

Best regards Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

On Friday 02 of March 2012 01:39:12 you wrote:
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} Email: hongwen.zhou-at-temple.edu Name: Hongwen Zhou
}
} Organization: Temple University
}
} Title-Subject: [Filtered] Images taken by CCD at low magnification inverted
}
} Message: The images were taken at magnifications of 3,000 and below at 80 kV. The electron-dense
} regions appear bright and the electron-transparent regions appear dark. The CCD gain reference
} images are up to date, although they were not taken at the same magnification. No such problems were
} observed at higher magnifications.
} Has anybody encountered the same problem? Any suggestions on how to correct it?
}
} Thanks!
}
} Hongwen Zhou
}
} ===========================
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} } Email: reganhll-at-gmail.com Name: Regan
} }
} } Organization: FMLS
} }
} } Title-Subject: [Filtered] 300KV Vs 100KV
} }
} } Message: Dear all,
} }
} } I was wondering what is the reason for the low contrast of images
} } in a 300KV Microscope compared to
} } an 100KV microscope. Technically the only difference is that you
} } use a beam of 0.00370nm for 100KV
} } and 0.00197nm for 300KV.
} } Theoretically if wavelength changes the resolution increases
} } However its unclear to me how it
} } affects contrast. there is a small indication however that this
} } affects mean free path by changing
} } the probability of scattering (p) defined as thickness /wavelength.
} } or something like that.
} }
} } I would be happy to some one explaining this elaborately.
} } Unfortunately I am biologist not a
} } physicist, But would be glad to see some equations if well
} } explained. or am I asking too much.
} }
} } Is it possible to have your 300 kv gun in F30 FEI microscope run in
} } 2 modes 300kv and 100kv?? Does
} } anybody have a microscope configured and using it in this way.
} } Greets, J
}
} Dear Regan,
} There are two types of contrast that are of concern to biologists,
} so I'll restrict my discussion to those. For stained, conventional
} plastic sections, the mechanism of contrast is that some electrons
} are scattered to large angles primarily by the high-Z atoms of the
} stain. These electrons are intercepted by the objective aperture
} (if no aperture is inserted, some electrons hit the objective lens
} lower pole piece, some are back-scattered, etc.), and these
} electrons are, therefore, removed from the image, creating dark
} regions. For a sufficiently thin specimen, fewer than half the
} electrons are scattered--check the pixel values of the dark vs.
} light regions of the image on a CCD camera--so the difference in
} contrast between 100 kV and 300 kV is due to the difference in
} scattering cross-section, with 100 kV electrons being scattered
} about twice as often as 300 kV electrons. For thicker or more
} heavily stained sections, this is not as much a factor, since even
} at 300 kV enough electrons are scattered to give contrast, and
} scattering from the unstained parts of the section at 100 kV
} decreases contrast by increasing the background.
} The second type of contrast occurs for unstained specimens such as
} frozen-hydrated ones. This is phase contrast, which is caused by
} the difference in phase change undergone by electrons traveling
} through a slab of ice compared to those that have traveled both
} through ice and biological material. The phase change is caused by
} the electrostatic potential within the specimen. This will decrease
} the wavelength of an electron traveling through a positive potential
} and increase the wavelength of an electron traveling through a
} negative potential. The relevant equations are E = T + V, or total
} energy (which is conserved) is equal to kinetic energy plus
} potential energy, V = qP, or potential energy equals the charge
} times the electrostatic potential, and v/lambda = ph, or the speed
} divided by the wavelength, which equals the circular frequency, is
} equal to the momentum divided by Plank's constant. (I think I got
} my factors of 2pi correct, but someone else from the list can
} correct me if I did not.). The changes in wavelength mean that the
} phase of each electron emerging from the specimen will depend on the
} potential it experiences along its path through the specimen. These
} differences are larger compared to 100 kV than to 300 kV, so for
} thin specimens, also referred to as weak phase objects, the contrast
} will be higher at 100 kV, all other things being equal. The
} principle other thing is the coherence of the beam. This has more
} of an effect on phase contrast than does accelerating voltage, and
} FEG sources are more coherent than LaB6 sources operated in "tip
} mode", which are, in turn, more coherent than LaB6 sources operated
} in the normal way with emission of electrons from both the tip and
} the pyramidal faces of the filament, or, least coherent, tungsten
} filaments.
} Imperfections in lenses, signal-to-noise ratio in biological
} images, and other factors such as the difference between the stain
} atoms and the biological features of interest, all have more
} influence on resolution than the theoretical maximum achievable due
} to wavelength. I hope this gives you a good introduction to
} contrast. There are any number of EM books that have several
} chapters each on contrast mechanisms and image formation. I suggest
} you find the one that most closely matches your knowledge of
} physics--which should not be completely lacking in a biologist,
} especially one who wants to see the equations.
} Finally, most scopes can be operated at several values of
} accelerating voltage. All that is necessary is to have good
} alignment files stored at the voltages of interest. (older, non-
} computerized scopes require realignment every time the HT is
} changed, but I assume you do not have one of those.) Every 300 kV
} instrument I have worked with has at least factory and installation
} alignments at 100 kV, so one can choose the operating voltage pretty
} freely.
} Yours,
} Bill
}
Dear List,
I guess that's what I get for posting on a Sunday and not considering
units. Plank's constant has units of momentum times distance, so
instead of v/lambda = ph, I should have had h=p*lambda.
Yours,
Bill

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Greetings Listers,
We are embarking on a project to fix, embed, and ultimately
prepare ultra-thin sections of a polysaccharide matrix. We are
considering the incorporation of ruthenium red in a typical fixative
such as osmium tetroxide or perhaps into another fixative, the use of a
fixative such as ruthenium tetroxide, and the use of durcupan as an
embedding medium. Durcupan might eliminate the shrinkage problem that
is associated with some other embedding media which require
dehydration. However, we understand that ultramicrotomy of
durcupan-embedded sections can be a challenge compared with conventional
media.

Would anyone out there be willing to share their experiences with
the above fixatives or with the embedding and ultramicrotomy of durcupan?

Thank you, Don

Donald Gantz
Dept. Physiology & Biophysics
Boston Univ. School of Medicine
700 Albany Street
Boston, MA 02118

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Hi,

We have been asked to cut thin sections of a carbon fiber reinforced resin
composite system. Question is as to whether we should risk our diamond
knives on these samples. this is a limited project so it does not pay us to
purchase any new knives for the samples. I would alike some input as to how
destructive this type of sample is to the knife edge.

Thanks in advance,

Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

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8, 28 -- From dsherman-at-purdue.edu Tue Mar 6 15:52:18 2012
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8, 28 -- Date: Tue, 6 Mar 2012 16:52:16 -0500
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Carlos;
I did my Ph.D. thesis analyzing steel samples, all highly ferromagnetic. The only time I had a problem was when the field of the lens pulled on the sample hard enough to overcome the snap ring (AKA Jesus Clip) and pulled the entire sample out of the holder. Well, that was a 100CX JEOL, and it was easy enough to disassemble and reassemble myself, and there was my sample hanging from the pole piece, all in one piece. Newer microscopes can be a lot more complex. I worked for 1.5 years at Komag division of Western Digital doing TEM of magnetic media. The total amount of magnetic material in those specimens was actually quite small and we never had any problems. I was told by the SEM folks, though, that over time, small grains can separate from full computer disks, especially if they were cut or scratched, or damaged during testing, and that these small particles would accumulate in the SEM final lens pole piece such that cleaning was required every few years. Apparently the biggest problem was in SEMS operated in immersion or snorkel mode.

John Mardinly, ASU
}
} ----------------------------------------------------------------------------
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}
} Dear colleagues,
}
} We do work with different kind of users that want to do TEM. Some of them =
} want to analyze magnetic material. Magnetic particles and bulk. Last week w=
} e had an accident with a sample. The sample was some king of iron alloy pre=
} pared by dimpler polishing. A large chunk of if broke inside the OL of our =
} TEM-FEG. Very fortunately that piece come out by itself. No need to call th=
} e service engineer to disassemble the OL and remove the broken sample.
}
} I was wondering how other EM labs deal with this kind of material. Do you =
} have some king of special protocol or just don't accept this kind of sample=
} inside the TEM?
}
} We have a service contract for the microscopes, but I did heard from the s=
} ervice engineer that using a magnetic material in a TEM is a misuse. And it=
} is not covered by our service contract. Some of my colleagues argues that =
} we can load any kind of sample in the TEM, and this should be covered by th=
} e contract. The fine line in the contract says it is a misuse as the engin=
} eer said. Any comment?
}
} What about magnetic particles? We have several users that study nanopartic=
} les. Magnetic or doped by magnetic elements like Co. Do you think these pa=
} rticle can fly off from the grid and get on the OL? These contamination on =
} the OL, can it affect the performance of the TEM?
}
} Thanks,
}
} Carlos
}
} __
} Carlos Kazuo Inoki
} Laboratório Nacional de Nanotecnologia
} Caixa Postal 6192
} Campinas, SP
} 13083-970
} Brazil
}
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Hi, Debby,

Try a couple of simple tests, if you would. First of all, I think you'll be O.K. with this project. You're not cutting diamond or glass. Trim your sample very small. Cut your fibers in cross section. Work with a small block face. Leave excess resin all around your sample. Try trimming the block with a razor blade. If the razor blade isn't chewed up when you trim a very thin slice off the block, this is a good sign. Then, if you have glass knives available, try cutting a few sections on one of these, or, preferrably, even an old diamond. I suspect that, since you're essentially cutting very dense carbon, you will be O.K.
I think your problem will be more with infiltration than with sectioning. I have no idea how you will get the fibers to infiltrate properly to allow you to keep them from turning to powder when you section them and when you put them in the scope. I would suggest using L. R. White, since it has the lowest viscosity of any resin, and use the softest formula you can, since the fibers will be very brittle to begin with. If this were my project, I would simply cut a very small block face of the sample on an old diamond knife, a knife where I knew I still had a part with some useable edge. I keep some old diamond knives around for cutting thick sections on, as these are much faster to use than glass knives, and I also use them for projects like this on occasion.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
________________________________________
X-from: dsherman-at-purdue.edu [dsherman-at-purdue.edu]
Sent: Tuesday, March 06, 2012 5:01 PM
To: Haller, Edward

Hi,

We have been asked to cut thin sections of a carbon fiber reinforced resin
composite system. Question is as to whether we should risk our diamond
knives on these samples. this is a limited project so it does not pay us to
purchase any new knives for the samples. I would alike some input as to how
destructive this type of sample is to the knife edge.

Thanks in advance,

Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

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Typically EDS detector resolution is reported as FWHM for MnKa.

In case spectra are acquired at MCA setting of 10eV per channel, then since
each channel is 10eV wide, there will be uncertainty of plus/minus 5eV in
defining resolution.

Here is an example:
If the FWHM falls between two channels, i.e., it is less than 14 channels
but more than 13 channels wide. Then how should resolution values be treated in such case?

If we plot the centroid value for each 10eV-wide channel and fit a curve
through the points then the width at half maximum with can be determined with
precision even better than 1eV.
Still such measurements do not change the fact that there is uncertainty of
10eV or plus/minus 5eV for any value of FWHM determined.

For example if resolution is determined as 135eV then due to the 10eV
channel width, such detector performance should be
defined as being better than 140eV and not worst than 130 eV, or if an
individual value is to be quoted then the plus/minus 5eV uncertainty
must be stated.

I am curious what the accepted agreement on this issue is and is there any
established standard procedure to determine and quote EDS detector resolution consistently.

Krassimir Bozhilov

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Is there any reason you couldn't cut them on glass and not risk the
diamonds?
I tend to use glass any time I am not sure of the sample.
David

}
} Hi,
}
} We have been asked to cut thin sections of a carbon fiber reinforced
} resin
} composite system. Question is as to whether we should risk our
} diamond
} knives on these samples. this is a limited project so it does not
} pay us to
} purchase any new knives for the samples. I would alike some input
} as to how
} destructive this type of sample is to the knife edge.
}
} Thanks in advance,
}
} Debby
}
} ---
} Debby Sherman, Interim Director Phone: 765-494-6666
} Life Science Microscopy Facility FAX: 765-494-5896
} Purdue University E-mail: dsherman-at-purdue.edu
} S-052 Whistler Building
} 170 S. University Street
} West Lafayette, IN 47907
} http://www.ag.purdue.edu/facilities/microscopy
}
}

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Dear list,
�
I am a little�bit lost with some definitions and would appreciate your help on this matter.
The main question is:
Is EDX (EDS, Energy dispersive spectroscopy) a part of XRF (X-ray fluorescence)?
�
When I visit the definition of XRF in wikipedia:
http://en.wikipedia.org/wiki/X-ray_fluorescence
�
I can read that "X-ray fluorescence (XRF) is the emission of characteristic "secondary" (or fluorescent) X-rays from a material that has been excited by bombarding with high-energy X-rays or gamma rays."
�
Since in TEM/SEM, the X-rays are produced by bombarding the specimen with electrons (and not x-rays or gamma rays as in the definition), it seems that EDS in TEM/SEM is not a part of XRF.
BUT later on the wiki page, I read that EDS is actually XRF.
�
I would be grateful for any comment.
�
regards,
Stephane

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2, 37 -- From: Stephane Nizet {nizets2-at-yahoo.com}
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Stephane

Wiki pages are not known for their technical accuracy, be careful when you use them.

X-ray Energy Dispersive Spectroscopy (XEDS) is a methodology used to measure a x-ray
emission from a material using technologies, which typically (but not exclusively) employ solid
state detectors and then graphically displays the result in the form of a pseudo-continuous
spectrum. The detectors used provide a signal which is proportional to the energy of the radiation
which is absorbed within their matrix. The signal is then amplified, processed and
displayed as a spectrum as a function of energy which created the signal in the detector.

There are a number of different detector technologies that can be used and include: Si(Li), HPGe,
SDD, HgI, Superconducting Transition Edge Microcalorimeters .....

XEDS has nothing to do with the excitation mechanism by which the x-rays are actually generated.

The x-rays can be created by any process, incident photons , electrons, ions ....

So saying that "EDS is actually XRF" is a misnomer. XRF is the signal excitation process, i.e.
x-ray emission from a specimen created by fluorescence (i.e. absorption of an x-ray and the
subsequent emission of x-ray photons of a different energy). This is different from the electron
column case which is an electron-in photon-out process. You can use any number of different
detectors to measure an XRF signal.

Of course,we are tacitly assuming that the article implies that XRF is being used to create x-rays!
Xray absorption can also create numerous other signals including electron emission this process is
called X-ray Photoelectron Spectroscopy (XPS) !

Cheers

Nestor
Your Friendly Neighborhood SysOp

On 3/8/12 7:24 AM, nizets2-at-yahoo.com wrote:
} ----------------------------------------------------------------------------
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}
} Dear list,
}
} I am a little bit lost with some definitions and would appreciate your help on this matter.
} The main question is:
} Is EDX (EDS, Energy dispersive spectroscopy) a part of XRF (X-ray fluorescence)?
}
} When I visit the definition of XRF in wikipedia:
} http://en.wikipedia.org/wiki/X-ray_fluorescence
}
} I can read that "X-ray fluorescence (XRF) is the emission of characteristic "secondary" (or fluorescent) X-rays from a material that has been excited by bombarding with high-energy X-rays or gamma rays."
}
} Since in TEM/SEM, the X-rays are produced by bombarding the specimen with electrons (and not x-rays or gamma rays as in the definition), it seems that EDS in TEM/SEM is not a part of XRF.
} BUT later on the wiki page, I read that EDS is actually XRF.
}
} I would be grateful for any comment.
}
} regards,
} Stephane
}
}
} ==============================Original Headers==============================
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--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory
Fellow of the Microscopy Society of America
E.P. Wigner Fellow - Oak Ridge National Laboratory
Senior Fellow of the Computational Institute - University of Chicago
Past-President Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a Mac !

===========================================

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Hi Amit,

This is interesting; I can think of a few things that may be going on.

One is that the 2100F is a 200kV field emission microscope and the F-30
is a 300kV machine and is probably FE as well, so there is a voltage
difference and the higher voltage machine will show less contrast from
the carbon film - but it is not a huge effect.

Another thing that I wonder about is carbon 'contamination' that can
wash-out contrast; I'm wondering whether you have a cold trap with
liquid nitrogen operating - to reduce contamination in one machine over
the other? Are you seeing contamination anyway?

Are you using the same aperture angles? If you have a large objective
aperture you will get less contrast from the carbon film - comparing
aperture angles between microscopes is easy if you use aluminum or gold
diffraction standards (used so you can check your camera length).

Have you tried STEM?

Good luck
Rob Keyse

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} Title-Subject: [Filtered] High background contrast
}
} Message: We are using JEOL JEM 2100F. But the contrast or diffraction from amorphous carbon from
} grid is too much. because of that we cannot sometimes see contrast from our samples clearly. same
} samples when done in Technai F-30 gives lot better contrast with back ground suppressed to almost
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--
Robert Keyse
EM Facility
Whitaker Laboratory
5 East Packer Avenue
Bethlehem
PA 18015
USA

Tel. +1 610 758 4283
Fax +1 610 758 4244

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Hi, Amit,

If I understand your question correctly, what I think you are asking is if you can enhance the appearance of the nanoparticles so you can see them better in the microscope. With certain particles, the answer would be, yes. It is possible to do silver or gold enhancement of metal nanoparticles to make the particles "grow" or become larger, chemically. This is a common procedure done in immunogold labeling. There are kits sold by the various electron microscope supply companies for this very procedure. The technique is fast and easy to do, normally done to gold particles. By varying time of exposure and/or temperature, you can end up with different size nanoparticles. Check with the vendors to see if this type of enhancement will work with the kind of nanoparticle that you are working with.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
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Email: amit.welcomes.u-at-gmail.com Name: Amit

Organization: Indian institute of science

Title-Subject: [Filtered] staining for inorganic particles

Message: I was just curious that like we have negative staining for nanoparticles coated with long
chain unsaturated molecules, biological samples etc, can we have some kind of staining for inorganic
shell of nano particles. eg if one metal is coated on another can we selectively bind some ligand to
outer shell etc to get better resolution? anyone tried such thing?

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Hi Stephane,
I think the alphabet soup is causing confusion between x-ray sources and
x-ray detection methods.

XRF (x-ray fluourescence) refers to the generation of characteristic x-rays
by a (relatively) monochromatic x-ray beam. The x-ray generation is very
good and with very little background. However, it is generally used for
bulk analysis because the beam is large and the penetration fairly deep.
There are micro-XRF systems now that can go inside an SEM chamber and
generate a fairly small spot, but still very large compared to an electron
beam.

EDS (energy dispersive spectrometry/spectrometer) refers to the means of
detecting the characteristic x-rays, whether they are generated by an x-ray
beam or an electron beam. WDS (wavelength dispersive
spectrometry/spectrometer) can also be used in either situation to detect
the characteristic x-rays that are generated.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Thursday, March 08, 2012 8:28 AM
To: kenconverse-at-qualityimages.biz

Dear list,
�
I am a little�bit lost with some definitions and would appreciate your help
on this matter.
The main question is:
Is EDX (EDS, Energy dispersive spectroscopy) a part of XRF (X-ray
fluorescence)?
�
When I visit the definition of XRF in wikipedia:
http://en.wikipedia.org/wiki/X-ray_fluorescence
�
I can read that "X-ray fluorescence (XRF) is the emission of characteristic
"secondary" (or fluorescent) X-rays from a material that has been excited by
bombarding with high-energy X-rays or gamma rays."
�
Since in TEM/SEM, the X-rays are produced by bombarding the specimen with
electrons (and not x-rays or gamma rays as in the definition), it seems that
EDS in TEM/SEM is not a part of XRF.
BUT later on the wiki page, I read that EDS is actually XRF.
�
I would be grateful for any comment.
�
regards,
Stephane

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Krassimir,
This is not my strong suit, but in looking at the results of software
calibration routines in modern EDS systems, I believe the bar graph that is
displayed, showing the actual output from the hardware, is integrated (from
my dim memory of calculus) into a smooth curve. If there are enough counts
to satisfy the statistical requirements (again, a dim math memory), then the
resulting calibration to the nearest 0.1eV centroid position is valid.

Using similar math, one could also determine the FWHM to a similar
precision.

I would love to know from those who are, in fact, good at this stuff if this
is the correct interpretation. If not, where have I gone wrong?

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Typically EDS detector resolution is reported as FWHM for MnKa.

In case spectra are acquired at MCA setting of 10eV per channel, then since
each channel is 10eV wide, there will be uncertainty of plus/minus 5eV in
defining resolution.

Here is an example:
If the FWHM falls between two channels, i.e., it is less than 14 channels
but more than 13 channels wide. Then how should resolution values be treated
in such case?

If we plot the centroid value for each 10eV-wide channel and fit a curve
through the points then the width at half maximum with can be determined
with
precision even better than 1eV.
Still such measurements do not change the fact that there is uncertainty of
10eV or plus/minus 5eV for any value of FWHM determined.

For example if resolution is determined as 135eV then due to the 10eV
channel width, such detector performance should be
defined as being better than 140eV and not worst than 130 eV, or if an
individual value is to be quoted then the plus/minus 5eV uncertainty
must be stated.

I am curious what the accepted agreement on this issue is and is there any
established standard procedure to determine and quote EDS detector
resolution consistently.

Krassimir Bozhilov

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Hello Amit,

You didn't say if you see the contrast on the negatives or on a digital
camera. If you are using a digital camera, please check your reference
images. If the reference images are not correct, they will add to the
background noise instead of suppressing it.

Mike Bode, Ph.D.
ResAlta Research Technologies Corp.

Mike.Bode-at-ResAltaTech.com
www.ResAltaTech.com

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Email: amit.welcomes.u-at-gmail.com Name: Amit

Organization: Indian institute of science

Title-Subject: [Filtered] High background contrast

Message: We are using JEOL JEM 2100F. But the contrast or diffraction from
amorphous carbon from grid is too much. because of that we cannot sometimes
see contrast from our samples clearly. same samples when done in Technai
F-30 gives lot better contrast with back ground suppressed to almost
invisibility! what can be the reason?

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Hello Amit,
My protocol in the JEM-2100F (with UHR pole piece [what do you have in the JEOL and what in the FEI microscopes?!]) for imaging tiny inorganic nanoparticles (smaller than 2 nm diameter) on strong contrasting (too thick) carbon film is this: the nanoparticles adsorbed to the supporting carbon film have to be presented at the top of the carbon film, when mounting the grid to the sample holder; like this the particles in the TEM will be localized above the carbon film, after inserting the sample holder; now focus on the carbon film so well, that the carbon film features will appear best resolved and therefore at lowest contrast (ideally the carbon film even becomes invisible); while the carbon film in the focus appears lowest contrasted, the nanoparticles on top of the carbon film are already slightly underfocussed und therefore can already produce some stronger contrasts in the image; maybe you want to underfocus just a little bit more, about 20 - 100 nm (at X100K), to gain a good compromise between well-focused-and-thus-low-contrasted-C-Film and already-underfocussed-and-therefore-better-contrasted-nanoparticles;
But the lighter the elements of the nanoparticles, the worth this protocol will be useful for you. If you want to image generally low contrasting specimens (i.e. proteins) than this will not work for you at all.
Best greetings from the EM-Labs of CIC biomaGUNE (SPAIN),
Marco M�ller

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Platform Manager - Electron Microscopy

CIC biomaGUNE
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20009 San Sebasti�n (Guip�zcoa)
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Email: amit.welcomes.u-at-gmail.com Name: Amit
Organization: Indian institute of science
Title-Subject: [Filtered] High background contrast

Message: We are using JEOL JEM 2100F. But the contrast or diffraction from amorphous carbon from grid is too much. because of that we cannot sometimes see contrast from our samples clearly. same samples when done in Technai F-30 gives lot better contrast with back ground suppressed to almost invisibility! what can be the reason?

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Hi, Amit,

PEG can be stained positively with uranyl acetate stain. In other words, it will stain with U.A. It is not a negative stain, but a positive stain. PEG will get dark when exposed to U.A. stain. The other alternative, as I mentioned, would be to grow the gold particles through silver or gold enhancement. Your silicone, magnesium and iron particles are already electron dense. I would be working at 60-80kV for this work, 100kV maximum, (I'm taking for granted that you're not trying for lattice resolution on anything here). Higher kV will not give you the contrast that you want. Use a 50 micron aperture for your work and you should have all the contrast you want.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
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X-from: Amit Gupta [amit.welcomes.u-at-gmail.com]
Sent: Thursday, March 08, 2012 10:56 AM
To: Haller, Edward

Hi, Amit,

If I understand your question correctly, what I think you are asking is if you can enhance the appearance of the nanoparticles so you can see them better in the microscope. With certain particles, the answer would be, yes. It is possible to do silver or gold enhancement of metal nanoparticles to make the particles "grow" or become larger, chemically. This is a common procedure done in immunogold labeling. There are kits sold by the various electron microscope supply companies for this very procedure. The technique is fast and easy to do, normally done to gold particles. By varying time of exposure and/or temperature, you can end up with different size nanoparticles. Check with the vendors to see if this type of enhancement will work with the kind of nanoparticle that you are working with.

Ed

Edward Haller, Lab Manager
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Organization: Indian institute of science

Title-Subject: [Filtered] staining for inorganic particles

Message: I was just curious that like we have negative staining for nanoparticles coated with long
chain unsaturated molecules, biological samples etc, can we have some kind of staining for inorganic
shell of nano particles. eg if one metal is coated on another can we selectively bind some ligand to
outer shell etc to get better resolution? anyone tried such thing?

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On Mar 8, 2012, at 5:43 AM, nizets2-at-yahoo.com wrote:

} I am a little bit lost with some definitions and would appreciate
} your help on this matter.
} The main question is:
} Is EDX (EDS, Energy dispersive spectroscopy) a part of XRF (X-ray
} fluorescence)?
}
} When I visit the definition of XRF in wikipedia:
} http://en.wikipedia.org/wiki/X-ray_fluorescence
}
} I can read that "X-ray fluorescence (XRF) is the emission of
} characteristic "secondary" (or fluorescent) X-rays from a material
} that has been excited by bombarding with high-energy X-rays or gamma
} rays."
}
} Since in TEM/SEM, the X-rays are produced by bombarding the specimen
} with electrons (and not x-rays or gamma rays as in the definition),
} it seems that EDS in TEM/SEM is not a part of XRF.
} BUT later on the wiki page, I read that EDS is actually XRF.
}
} I would be grateful for any comment.
}
Dear Stephane,
Another way to say it is that XRF is x-ray induced x-ray emission and
EDS is electron induced x-ray emission. There is also particle
induced x-ray emission, which usually refers to the use of protons or
alpha particles to produce characteristic x-rays. These are obviously
closely related, and one could call them all parts of the same
technique, but I would not call them the same because different
instruments are used to perform them. Wikipedia is very useful, but I
am not a fundamentalist, so I must admit that there is always the
possibility of mistakes is any literature.
Yours,
Bill

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To all,

Our lab has a couple of whole slide scanners which are somewhat sensitive to vibrations caused by users tapping/bumping the tables on which the scanners rest. �For clarity, the computers controlling the scanning process are located adjacent to the scanners on the same table. �We've tried resting a scanner directly on anti-vibration pads (rubber or rubber with cork core) but the equipment is too lightweight (55 Lbs, 25 kg) to benefit. �In the near future we will be relocating our lab, and the equipment will be moving from tables to standard laboratory benches. �I'm hoping the lab benches will be less sensitive to user-induced vibration, but not holding my breath!

Has anyone had success with a passive desktop anti-vibration system designed to isolate just one light-weight piece of equipment, not the entire table-top? �Active pneumatic systems would be price-prohibitive, and likely overkill.�

Thank you for your suggestions!

Karen Zaruba

University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Ed,
I take advantage of your post for asking you some help: I'm looking for
literature on PEG staining at this very moment, and your post looked
like sent by Heavens :)
I've found some papers about PEG staining with U.A., but none specifies
staining procedure details (e.g. "exposure" time to U.A.), and I've
never done any staining at all. Besides, in our lab we've already Uranyl
Nitrate: I guess it should work as well, but haven't found any hint
about it.
Could you please suggest me any text or give me some recommendation?
If you think this is off topic and prefer to keep the thread in topic,
you could answer me directly.
Thanks!

Davide

~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~
Davide Cristofori

direct phone = 041.234.67.26
e-mail = dcristofori[at]unive[dot]it

Universita' Ca' Foscari Venezia
Dipartimento di Sc. Molecolari e Nanosistemi
Lab. di Scienza e Tecnologia dei Materiali
Via Torino, 155b
I-30172 Mestre (VE)
Italy
~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~

Il 08/03/2012 18.02, ehaller-at-health.usf.edu ha scritto:
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} Hi, Amit,
}
} PEG can be stained positively with uranyl acetate stain. In other words, it will stain with U.A. It is not a negative stain, but a positive stain. PEG will get dark when exposed to U.A. stain. The other alternative, as I mentioned, would be to grow the gold particles through silver or gold enhancement. Your silicone, magnesium and iron particles are already electron dense. I would be working at 60-80kV for this work, 100kV maximum, (I'm taking for granted that you're not trying for lattice resolution on anything here). Higher kV will not give you the contrast that you want. Use a 50 micron aperture for your work and you should have all the contrast you want.
}
} Ed
}
} Edward Haller, Lab Manager
} University of South Florida
} Integrative Biology Department
} Electron Microscopy Core
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} ehaller-at-usf.edu
} Office: ISA 1046
} ________________________________________
} X-from: Amit Gupta [amit.welcomes.u-at-gmail.com]
} Sent: Thursday, March 08, 2012 10:56 AM
} To: Haller, Edward
} Subject: Re: [Microscopy] RE: viaWWW:staining for inorganic particles
}
} what i want is suppose i have a core-shell particle. an inorganic core say silicon carbide and over it is thin shell of SiGe or Fe2O3 with shell of MgO etc etc. then can i enhance the contrast between the chemical moieties? like i read one paper where the group was doing the same with negative staining of gold nano particle coated with (if i remember correctly) PEG.
}
} On Thu, Mar 8, 2012 at 7:52 PM, {ehaller-at-health.usf.edu {mailto:ehaller-at-health.usf.edu} } wrote:
}
}
}
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} Hi, Amit,
}
} If I understand your question correctly, what I think you are asking is if you can enhance the appearance of the nanoparticles so you can see them better in the microscope. With certain particles, the answer would be, yes. It is possible to do silver or gold enhancement of metal nanoparticles to make the particles "grow" or become larger, chemically. This is a common procedure done in immunogold labeling. There are kits sold by the various electron microscope supply companies for this very procedure. The technique is fast and easy to do, normally done to gold particles. By varying time of exposure and/or temperature, you can end up with different size nanoparticles. Check with the vendors to see if this type of enhancement will work with the kind of nanoparticle that you are working with.
}
} Ed
}
} Edward Haller, Lab Manager
} University of South Florida
} Integrative Biology Department
} Electron Microscopy Core
} SCA 110
} 4202 East Fowler Avenue
} Tampa, FL 33620
} 813-974-2676 {tel:813-974-2676}
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} Email: amit.welcomes.u-at-gmail.com {mailto:amit.welcomes.u-at-gmail.com} Name: Amit
}
} Organization: Indian institute of science
}
} Title-Subject: [Filtered] staining for inorganic particles
}
} Message: I was just curious that like we have negative staining for nanoparticles coated with long
} chain unsaturated molecules, biological samples etc, can we have some kind of staining for inorganic
} shell of nano particles. eg if one metal is coated on another can we selectively bind some ligand to
} outer shell etc to get better resolution? anyone tried such thing?
}
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7, 21 -- From dcristofori-at-unive.it Thu Mar 8 11:57:26 2012
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On Mar 8, 2012, at 5:43 AM, microscopylistserver-
noreply-at-microscopy.com wrote:

} Email: amit.welcomes.u-at-gmail.com Name: Amit
}
} Organization: Indian institute of science
}
} Title-Subject: [Filtered] staining for inorganic particles
}
} Message: I was just curious that like we have negative staining for
} nanoparticles coated with long
} chain unsaturated molecules, biological samples etc, can we have
} some kind of staining for inorganic
} shell of nano particles. eg if one metal is coated on another can we
} selectively bind some ligand to
} outer shell etc to get better resolution? anyone tried such thing?
}
Dear Amit,
Another possibility no one else has yet mentioned is to evaporate a
metal onto the specimen; e.g., platinum shadowing. If the specimen is
high-Z, it will be difficult to distinguish the shadow from the shell,
but energy filtering could do that. In any case, the resolution is
unlikely to be improved by staining, since it will be limited to the
size of the stain particles. There are high-resolution techniques for
depositing a layer of Cr that are used in HRSEM, so maybe you could
try something like that.
Yours,
Bill

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Thanks to all who replied so quickly to my question! �

Most of the responses have involved using a marble, granite, steel or other heavy slab to add weight and stability above a shock absorber of some type. �I like best the idea of putting to use those dampening pads I already have (affectionately known around the lab as "scotcheroos" after the butterscotch bars they resemble). �Following one suggestion I am searching our campus surplus for a slab. �Possibly I will have a very cost-effective solution in the near future.

However, I would still welcome suggestions beyond the realm of granite slabs, in the event we do find a small outlay of funds within the budget. �Also the scotcheroos may not cut the mustard, so to speak.

Sincerely,

Karen

Karen Zaruba
University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Karen:

Marble or granite balance tables are not cheap, one can cost over $1000.
Just google "marble balance table"
They are the best option I know of short of an pneumatic antivibration
table.put the scotcheroos between the floor and the legs and between the
legs and the table top

Bob
Bob
Dr. Robert J. Schmitz
Associate Professor of Anatomical Sciences
Department of Biology
University of Wisconsin-Stevens Point
800 Reserve Street, 380 TNR Bld
Stevens Point, WI 54481
715-346-2420
Email: rschmitz-at-uwsp.edu
http://www4.uwsp.edu/biology/faculty/RSchmitz/

On 3/8/12 3:30 PM, "kstzar-at-yahoo.com" {kstzar-at-yahoo.com} wrote:

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If you need only a slab of stone (as opposed to a whole table) you
can get a slab of granite or other stone from a local quarry or mason
for a lot less than $1000. The finish would be not so fancy. Probably
doesnt matter as long as it is reasonably flat on one surface.
Likewise, depending on how much engineering/physics is done around
your University you might be able to scrounge a large plate of steel.

Tobias

}
} Karen:
}
} Marble or granite balance tables are not cheap, one can cost over $1000.
} Just google "marble balance table"
} They are the best option I know of short of an pneumatic antivibration
} table.put the scotcheroos between the floor and the legs and between the
} legs and the table top
}
}
} Bob
} Bob
} Dr. Robert J. Schmitz
} Associate Professor of Anatomical Sciences
} Department of Biology
} University of Wisconsin-Stevens Point
} 800 Reserve Street, 380 TNR Bld
} Stevens Point, WI 54481
} 715-346-2420
} Email: rschmitz-at-uwsp.edu
} http://www4.uwsp.edu/biology/faculty/RSchmitz/
}
}
}
}
} On 3/8/12 3:30 PM, "kstzar-at-yahoo.com" {kstzar-at-yahoo.com} wrote:
}
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--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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You could buy a bag of concrete mix at your local building supply and make up a slab to your liking. You could make the slab any size or weight that you would like. If you know the characteristics of your isolation blocks, you could design the weight so the resonance is far removed from the environmental vibrations.

The civil engineering department at U of M should have the skills to build the forms, mix the concrete, and finish it off nicely. It could be practical experience for the students.

You could also probably find someone in the mechanical engineering of engineering mechanics departments to help with the vibrational analysis. They should have the equipment to characterize your isolation pads and maybe characterize the room - or the students tapping on the table.

I would even think both departments would appreciate a real-world application of their disciplines.

Warren

-----Original Message-----
X-from: kstzar-at-yahoo.com [mailto:kstzar-at-yahoo.com]
Sent: Thursday, March 08, 2012 3:30 PM
To: wesaia-at-iastate.edu

Thanks to all who replied so quickly to my question! �

Most of the responses have involved using a marble, granite, steel or other heavy slab to add weight and stability above a shock absorber of some type. �I like best the idea of putting to use those dampening pads I already have (affectionately known around the lab as "scotcheroos" after the butterscotch bars they resemble). �Following one suggestion I am searching our campus surplus for a slab. �Possibly I will have a very cost-effective solution in the near future.

However, I would still welcome suggestions beyond the realm of granite slabs, in the event we do find a small outlay of funds within the budget. �Also the scotcheroos may not cut the mustard, so to speak.

Sincerely,

Karen

Karen Zaruba
University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Hi Amit
If you 2100F has STEM try dark field for imaging small particles-they will jump out at you that way.

In TEM mode check the auto contrast limits on the digital images, use a small objective aperture and
try lower KV. What CCD camera and software are you using? If Gatan DM change the gamma setting on
the image to enhance the particle contrast.

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

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} Title-Subject: [Filtered] High background contrast
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Email: mark.auty-at-teagasc.ie Name: Mark Auty

Organization: Teagasc

Title-Subject: [Filtered] PhD opportunity - food microstructure

Message: See below PhD studentship on food microstructure:

Teagasc Walsh Fellowship Opportunity

The development of nano-engineered SMART dairy ingredients with enhanced process stability (Ref:
2012204)

Background

Teagasc Food Research Centre, part of Teagasc the Irish Agriculture and Food Development
Authority, undertakes public scientific research and provides technological services to the food
industry with emphasis on functional foods, dairy, food ingredients and formulated foods and
beverages. The programme consists of fundamental and applied research, technology transfer,
industrial consultancy and training under the overall umbrella of an innovation management strategy.
The Food Chemistry and Technology Department at Teagasc is primarily responsible for innovation and
research in the dairy processing sector nationally, and is actively working with all major
ingredient suppliers and infant milk formulae manufacturers based in Ireland. It is currently the
only research provider in Ireland which supports research on dairy proteins and related separations
and spray drying technologies. To this end, Teagasc has obtained funding from the Dairy Levy for a
new project: MDDT-0103-6261 SMART milk ingredients. The overall aim of this project is to develop
key scientific capability in the area of protein chemistry by exploiting the protein-based
aggregation kinetics in milk and to add value to milk based ingredients for use in products for
export. The project will build a knowledge base for protein structure-functionality and expand its ro

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Dear List
We need to purchase membrane filters, to place on them blood and other cells
and examine with SEM. As I see in the market, track etched filters have
single pores that make a much better background. There are two types
avalable, Polycarbonate (PCTE) and Polyester (PETE) track etched membranes
with identical shapes, yet PETE is more resistant to solvents than PCTE. If
anybody has used both, I would like to ask if there is any difference in SEM
imaging due to possible differences in electron conductivity or other
properties that may affect the quality of imaging.
Thanks in advance
yorgos

Dr Yorgos Nikas
Athens Innovative Microscopy
Skra 36 Voula 16673 GREECE

Tel/fax +30 210 8957677
mobile +30 6945 107477
eikonika-at-otenet.gr
www.aim.cat
***********************************

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We have been using polycarbonate filters, 12mm diameter, 0,2micron for the preparation of just about all kinds of cells.
The results were always great.
Recently we started seeing crystals on the filters in the SEM.
After much trial and error we read the notice on the manufacturer's packaging stating that a change has been made to the product.
Apparently some kind of coating is now put on the filter and this, we think, crystalizes with dehydration or CPD.
Anyone with a similar experience?
Alan Hall

Laboratory for Microscopy & Microanalysis
Room1-33, Natural Sciences 2
University of Pretoria
Lynnwood Road
Pretoria 0002
South Africa
Tel: +27 12 4202075
Fax: +27 12 3625150
Mobile: +27 82 3319040

Laboratory for Microscopy & Microanalysis
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Hatfield 0028
South Africa

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Dear List
We need to purchase membrane filters, to place on them blood and other cells
and examine with SEM. As I see in the market, track etched filters have
single pores that make a much better background. There are two types
avalable, Polycarbonate (PCTE) and Polyester (PETE) track etched membranes
with identical shapes, yet PETE is more resistant to solvents than PCTE. If
anybody has used both, I would like to ask if there is any difference in SEM
imaging due to possible differences in electron conductivity or other
properties that may affect the quality of imaging.
Thanks in advance
yorgos

Dr Yorgos Nikas
Athens Innovative Microscopy
Skra 36 Voula 16673 GREECE

Tel/fax +30 210 8957677
mobile +30 6945 107477
eikonika-at-otenet.gr
www.aim.cat
***********************************

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} } }
} We have been using polycarbonate filters, 12mm diameter, 0,2micron for the
} preparation of just about all kinds of cells.
} The results were always great.
} Recently we started seeing crystals on the filters in the SEM.
} After much trial and error we read the notice on the manufacturer's
} packaging stating that a change has been made to the product.
} Apparently some kind of coating is now put on the filter and this, we think,
} crystalizes with dehydration or CPD.
} Anyone with a similar experience?

No, fortunately not, so far.
But from my point of view, it makes sense to confront the manufacturer with your results (send them an example SEM image BEFORE and AFTER change of the product!) and ask for a statement, or improvement of the product, the filter ...
kind regards,
Reinhard

--
Prof. Dr. Reinhard Rachel
University of Regensburg
Centre for EM / Anatomy
Faculty of Biology & Preclin. Med.
Universitaetsstrasse 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-regensburg.de
office: VKL 3.1.29

next microscopy conferences:
http://www.emc2012.org.uk/
EMC 2012 in Manchester, UK
http://www.mc2013.de/
MC2013 in Regensburg, Germany

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7, 25 -- To: "microscopy listserver" {Microscopy-at-microscopy.com} , {Alan.Hall-at-up.ac.za}
7, 25 -- Subject: membrane filters in SEM
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Dear All,
maybe someone has tried to image fine particles which needed to be
critical point dried before going into the SEM.
Like dust particles which also have organic particles with them.
Without drying or a lot of sputtercoating (which is prohibitve because I
need to image at least at ca. 10nm resolution) I got charge/discharge
effects.
Any idea how to fix the particles on the substrate before going intop
CPD? And still have some remaining after CPD?
I thought about trying Polysine coated coverslips or something other
"sticky".
But I also need o smooth background surface...

Best wishes,
Stefan

--

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Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
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Anfahrt: http://Mail.map24.com/Stefan.Diller
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Hi, Karen,

For years, what we used to mount ultramicrotomes on were stacks of concrete cinderblocks topped with "I" shaped concrete capstones, all purchased from either Home Depot or Lowes. The capstones are monsterously heavy, requiring several grown men to lift them safely. You might want to consider going this route. It's far less expensive than any other route. You can top the homemade table off with a wooden top isolated from the concrete by your vibration "scotcheroos" to dress the table up a little bit and give you a smooth working surface.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
________________________________________
X-from: kstzar-at-yahoo.com [kstzar-at-yahoo.com]
Sent: Thursday, March 08, 2012 4:36 PM
To: Haller, Edward

Thanks to all who replied so quickly to my question!

Most of the responses have involved using a marble, granite, steel or other heavy slab to add weight and stability above a shock absorber of some type. I like best the idea of putting to use those dampening pads I already have (affectionately known around the lab as "scotcheroos" after the butterscotch bars they resemble). Following one suggestion I am searching our campus surplus for a slab. Possibly I will have a very cost-effective solution in the near future.

However, I would still welcome suggestions beyond the realm of granite slabs, in the event we do find a small outlay of funds within the budget. Also the scotcheroos may not cut the mustard, so to speak.

Sincerely,

Karen

Karen Zaruba
University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Sorry if I'm repeating someone (I just caught the end of this thread)
but years ago we had a very nice granite microtome table made to our
specifications by a local tombstone company for a very reasonable price
(I think a few hundred dollars). They had all the tools and hardware
necessary to connect the pieces together, and the cranes and dollies to
deliver it and place it where we wanted. I think it was probably a tax
write-off for them, so if you approach them from this angle, you might
be able to land a bargain.

Jim

--

James M. Ehrman
Digital Microscopy Facility
Mount Allison University
63B York St.
Sackville, NB E4L 1G7
CANADA

phone: 506-364-2519
fax: 506-364-2505
email: jehrman-at-mta.ca
www: http://www.mta.ca/dmf

Clones are people two.

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Hi Karen,

If your slide scanner was more like my Epson V750 Pro 35mm film type scanner
[weighing 15lbs], I use a 1/2" marble chopping board from a supermarket that
cost �8 and added in two superbouncy 3.5cm solid rubber balls cut in half to
give four feet [flat side to table top, and the base of the half ball glued
to thick felt]. This works well enough in the home office, and you can go
upmarket and buy a classier �30 marble chopping board. At home I mount my
desktop tower PC case on the same half-bouncyballs as well to minimise
vibration to the hard drives etc.

However your scanner, at 55lb, seems in need of more serious anti-vibration,
so I'd check out a thick slate slab as well [as Guy mentioned last year, its
often far cheaper than granite to get cut to size and delivered. Plus slate
looks/feels a lot nicer than a concrete]. To support a heavy slab you need
more serious antivibration pads and these are available commercially from
many sources. We always used Fabreeka Fabcel pads
[http://www.fabreeka.com/documents/file/products/FabcelPad.pdf], a sort of
honeycombed soft nitrile rubber mat that can be cut up into smaller pads
[say 4x4 inches] and multiple stacked one pad on another to improve their
effectiveness [stuck together with double sided tape]. Fabcel pads come in a
variety of grades [softness] depending on the weight supported - we probably
used the lightest type 25. Although we used massive 110lb mild steel slabs
in the past under microscopes as they were so much cheaper than granite,
they did need covering in Fablon to look nice and they rang like a bell if
hit which isn't ideal [but they did work well enough when used with the
lighter grade Fabcel pads and a thin ribbed rubber mat glued to the top].
The Fabreeka pads are often strangely covered in a thin layer of silicon
oil, which can be wiped off [mostly], but they sit well out of the way under
the heavy slab.

Also consider reinforcing the lab worktop from underneath with extra support
legs, if the worktop is thin and flimsy - this extra rigidity under the area
of the anti-vibrational slab really helped us and cost around �100 for a 4
leg frame with adjustable feet via our workshop.

Regards

Keith

---------------------------------------------------------------------------
Dr Keith J. Morris,
Molecular Cytogenetics and Microscopy Core,
Laboratory 00/069 and 00/070,
The Wellcome Trust Centre for Human Genetics,
Roosevelt Drive,
Oxford� OX3 7BN,
United Kingdom.

Telephone:� +44 (0)1865 287568
Email:� kjmorris-at-well.ox.ac.uk
Web-pages: http://www.well.ox.ac.uk/molecular-cytogenetics-and-microscopy

-----Original Message-----
X-from: kstzar-at-yahoo.com [mailto:kstzar-at-yahoo.com]
Sent: 08 March 2012 17:46
To: kjmorris-at-well.ox.ac.uk

To all,

Our lab has a couple of whole slide scanners which are somewhat sensitive to
vibrations caused by users tapping/bumping the tables on which the scanners
rest. �For clarity, the computers controlling the scanning process are
located adjacent to the scanners on the same table. �We've tried resting a
scanner directly on anti-vibration pads (rubber or rubber with cork core)
but the equipment is too lightweight (55 Lbs, 25 kg) to benefit. �In the
near future we will be relocating our lab, and the equipment will be moving
from tables to standard laboratory benches. �I'm hoping the lab benches will
be less sensitive to user-induced vibration, but not holding my breath!

Has anyone had success with a passive desktop anti-vibration system designed
to isolate just one light-weight piece of equipment, not the entire
table-top? �Active pneumatic systems would be price-prohibitive, and likely
overkill.�

Thank you for your suggestions!

Karen Zaruba

University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Karen,

Ed's description below is the type of concrete top that I had written to you about mounted on 3 legs of cinderblocks. For others who did not see my message, the table should not touch the wall or other cabinets. The contractors who built my table the first time were not too pleased that they needed to take sledge hammers to the support legs to move it two inches away from the wall!

I used a pretty vinyl table cloth folded in half to cover the top and front of my table (used with ultra microtomes on a rubber pad). It lasted about 10 years and saved my forearms from getting scratched up - besides brightening up my lab.

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E - Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov

Opinions and experiences related are those of Pat Connelly and do not represent the NIH. This message is not confidential and can be freely shared and reproduced.

__________
Hi, Karen,

For years, what we used to mount ultramicrotomes on were stacks of concrete cinderblocks topped with "I" shaped concrete capstones, all purchased from either Home Depot or Lowes. The capstones are monsterously heavy, requiring several grown men to lift them safely. You might want to consider going this route. It's far less expensive than any other route. You can top the homemade table off with a wooden top isolated from the concrete by your vibration "scotcheroos" to dress the table up a little bit and give you a smooth working surface.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
________________________________________
X-from: kstzar-at-yahoo.com [kstzar-at-yahoo.com]

Thanks to all who replied so quickly to my question!

Most of the responses have involved using a marble, granite, steel or other heavy slab to add weight and stability above a shock absorber of some type. I like best the idea of putting to use those dampening pads I already have (affectionately known around the lab as "scotcheroos" after the butterscotch bars they resemble). Following one suggestion I am searching our campus surplus for a slab. Possibly I will have a very cost-effective solution in the near future.

However, I would still welcome suggestions beyond the realm of granite slabs, in the event we do find a small outlay of funds within the budget. Also the scotcheroos may not cut the mustard, so to speak.

Sincerely,
Karen

Karen Zaruba
University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Hi Stefan,
Have you considered using TEM grids with functionalized surfaces? The
functionalized surfaces have positive or negative charge or other chemical
functionality and behave similarly to "adhesive" slides. Particles anchored
on the surface of a functionalized grid should stick throughout CPD. Also,
the silicon grid form provides a very uniform background. These can be
purchased through Plano GmbH. You can read about the variety of
functionalizations available at
http://www.dunesciences.com/biogrids.html.

John Miller, Ph.D.
Dune Sciences, Inc.
Phone: 541-359-4710
jmiller-at-dunesciences.com
www.dunesciences.com

-----Original Message-----
X-from: stefan.diller-at-t-online.de [mailto:stefan.diller-at-t-online.de]
Sent: Friday, March 09, 2012 4:22 AM
To: jmiller-at-dunesciences.com

Dear All,
maybe someone has tried to image fine particles which needed to be critical
point dried before going into the SEM.
Like dust particles which also have organic particles with them.
Without drying or a lot of sputtercoating (which is prohibitve because I
need to image at least at ca. 10nm resolution) I got charge/discharge
effects.
Any idea how to fix the particles on the substrate before going intop CPD?
And still have some remaining after CPD?
I thought about trying Polysine coated coverslips or something other
"sticky".
But I also need o smooth background surface...

Best wishes,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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Dear List Members,

Electron Microscopy Laboratory of the Physics Department, Aristotle
University of Thessaloniki, Greece, is organizing a Summer School on

Micro- and Nano- structural characterization of materials, focused on
electron microscopy

on July 11-15, 2012

The school is addressed to PhD and postgraduate students, as well as all
young researchers interested in structural characterization and electron
microscopy. Apart from the lectures, a hands-on training on specimen
preparation techniques will also be offered.

For more information, please visit:
http://micronano2012.physics.auth.gr

Best regards,
E.K. Polychroniadis

Prof. E.K. Polychroniadis
Department of Physics
Aristotle University of Thessaloniki
Thessaloniki 54124, Greece
Tel.: +30.2310.998163
Fax: +30.2310.998241
e-mail: polychr-at-auth.gr

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Dear list
I have been asked to examine some blastocysts in the esem. The cells will be in a suspension and will be very low in number. I have imaged lots of cultured cells but always cultured on a substrate. Could some of you please advise on the best way to handle these cells for esem. Could I simply put a drop of suspension onto a coverslip or would using some sort of filter be better? The researcher wants to use esem to look at the cells in wet mode.
Best wishes
Nicola

Mrs Nicola Weston
Electron Microscope Technician
Lab Rm 103/Office Rm 109
Wolfson Building
School M3
University of Nottingham
Nottingham NG7 2RD
Tel: +44 (0) 115 951 3759

This message and any attachment are intended solely for the addressee and may contain confidential information. If you have received this message in error, please send it back to me, and immediately delete it. Please do not use, copy or disclose the information contained in this message or in any attachment. Any views or opinions expressed by the author of this email do not necessarily reflect the views of the University of Nottingham.

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Dear Karen,
Minus-K Technology manufactures a compact, passive vibration isolation
system that is used for small atomic force microscopes. It could well suit
your application. We have used units available at a discount price.
Please contact me or jon-at-asmicro.com offline to discuss the size of your
equipment, the size of the Minus-K unit and the price.

Disclaimer: We are not affiliated with Minus-K Technology. We do buy,
refurbish, sell and install used NanoScope Atomic force and scanning probe
microscopes. Some of the AFM systems we have bought include the Minus-K
antivibration unit, and we have some in stock now.

regards,
Don
=============================================
Don Chernoff, Ph.D., President
Advanced Surface Microscopy, Inc. E-Mail: donc-at-asmicro.com
3250 N. Post Rd., Ste. 120 Voice: 317-895-5630
INDIANAPOLIS IN 46226 USA Toll free: 800-374-8557 (in USA & Canada)
web: http://www.asmicro.com Fax: 317-895-5652
[business activities: analytical services in AFM, AFM probes, consulting,
training,
calibration and test specimens, calibration and measurement software,
used NanoScope equipment.] 2009-2010 is our 20th year in business!
=============================================
----- Original Message -----
From: kstzar-at-yahoo.com
To: donc-at-asmicro.com
Sent: Thursday, March 08, 2012 1:43 PM
Subject: [a] [Microscopy] LM: Passive desktop anti-vibration solutions

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To all,

Our lab has a couple of whole slide scanners which are somewhat sensitive
to vibrations caused by users tapping/bumping the tables on which the
scanners rest. For clarity, the computers controlling the scanning process
are located adjacent to the scanners on the same table. We've tried resting
a scanner directly on anti-vibration pads (rubber or rubber with cork core)
but the equipment is too lightweight (55 Lbs, 25 kg) to benefit. In the
near future we will be relocating our lab, and the equipment will be moving
from tables to standard laboratory benches. I'm hoping the lab benches will
be less sensitive to user-induced vibration, but not holding my breath!

Has anyone had success with a passive desktop anti-vibration system
designed to isolate just one light-weight piece of equipment, not the entire
table-top? Active pneumatic systems would be price-prohibitive, and likely
overkill.

Thank you for your suggestions!

Karen Zaruba

University of Minnesota
Division of Infectious Diseases
Department of Medicine
A638 Mayo Building
420 Delaware St. SE
Minneapolis, MN 55455

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Before I begin, you should be using the Mag-i-cal sample for calibration.
You can calibrate all of the TEM measurements with that one sample. For
magnification purposes, you can go from very low mag to very high mag and
you don't need to have two samples. It also can be used for calibrating the
camera constant for diffraction and the diffraction pattern rotation
relative to the image. You also don't need to have a NIST traceable
standard, since it is referenced to the lattice spacing of silicon. Of
course, you will have to re-write your calibration operating procedure to
reflect that.

I used both samples in my early days at PPG on a 120 keV machine. I noticed
that the spheres could shrink when there was only one or two in the field of
view and I viewed them for any period of time. On a 200 keV machine, this
was less of a problem. I minimized the problem by shifting the sample into
the field of view and immediately collecting an image. The best way to do
the diffraction grating and spheres is to have them in the same sample.
That way you have the cross over point(s) where you can correlate the two
calibrations with both standards. I'm sure that you will find that they
don't agree, as did I. (The Mag-i-cal sample uses the Si d-spacings as its
internal standard for the rest of the distances on the sample. In other
words, its traceable back to the fundamental lattice spacing of Si. )

You will find that the microscope, even when set up perfectly, can have the
magnification off by as much as 5-10%. With respect to your QC guy, who
cares? What is more important is reproducibility. If you redo the sample
several times after taking it out of the microscope and putting it back in
and redoing the eucentric position and are getting the same results, you're
golden. Since you're doing the calibration, probably on a routine basis,
as specified by your standard operating procedures for calibration, just
follow those procedures and he can't really say anything. If you want to
change your procedures, then you have to rewrite those. If you have set up
the sample in the eucentric height and collected the image at focus, measure
the mag from your standard, find the "true" magnification with the error,
and record the correction for each magnification. For my error reporting, I
would take several images at each mag and double the standard deviation of
the measurements. Record those corrections and apply them to your images
according to your procedures. Don't let him talk you into getting the
manufacturer out to work on the microscope to try to fix it. Just wait for
the next maintenance call, tell the service engineer and show him/her the
results. Afterwards, just redo the calibration series as you need to do if
they do anything major to the instrument.

For digital images, all you really want is the nm/pixel value for each
magnification setting, assuming that you always set up the digital camera
the same way. The camera resolution is not going to affect your measurement
unless the known feature size in your standard is on the order of a few
pixels. So, just don't do that.

If you are getting reproducible values for the corrections, you are doing
the calibration properly. You will probably also see a similar
reproducible error between the two standards at the magnifications where you
can use both.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: frank_karl-at-ardl.com [mailto:frank_karl-at-ardl.com]
Sent: Thursday, March 01, 2012 10:24 AM
To: s.walck-at-cox.net

Let me ask a couple of questions about calibrating a TEM, or rather
verification of calibration.

I calibrate with a diffraction grating replica at low magnification and
follow the procedure outlined by Ernest Fullam: I measure the same particle
at= several calibrated magnification and use an average value to calibrate
my = higher magnification.
So far so good!

Then I use NIST traceable Nanospheres to verify the calibration.

I'm in discussions with our QC guy. He a nice fellow and we both want to
improve the measurement values we collect. Unfortunately, my nanosphere
average values aren't falling close enough to the publish value.

I'm sure the scope is set up properly and the sample height is correct.

So, my two questions are,
Do other TEM operators verify their magnifications and what are do you
consider an acceptable difference? 1%, 8%, one standard deviations or
??????
And
Since the camera seems to be the connecting bridge between the image and
measurement, does anyone know of literature or have experience with camera
resolution and its affect on measurements?

Thanks for your help.................

Frank

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26, 42 -- Subject: RE: [Microscopy] Verification and the QC guy
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Contents Retrieved from Microscopy Listserver Archives
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Good Morning

I know there has been some discussion in the past regarding the life expectancy of instruments with consensus running at between 20-40 years. I would like to ask what is the general practice for writing down the asset value? Would you consider the Asset Life to be 10 years? Would the same apply to other equipment such as ultratomes, or CCD cameras?

Thanks in advance
Naomi

Naomi McCallum

Pathology Queensland
Australia

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7, 25 -- From: "Naomi McCallum" {Naomi_McCallum-at-health.qld.gov.au}
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Registration and hotel lines are open; the abstract deadline has past,
and hopefully you are making plans to attend the Microscopy and
Microanalysis (M&M) 2012 meetings in Phoenix, July 29-Aug. 2.

For all students (or those of you that may have students) planning to
attend these meetings, please consider the student bursary program
offered by MSA. The purpose of this program is to encourage students
to
attend the annual meeting, where they can meet and interact with the
established microscopy community while at the same time defraying some
meeting costs.

The students work for ~20 hours (or up to 40 hours) during the meeting
or pre-meeting events and are paid $10 an hour. The jobs involve such
things as providing support in the different symposia (assisting with
audio-visual needs, speaker set-up, maintaining an attendance count),
staffing the volunteer office, monitoring use of the Internet Caf�,
and
helping with vendor tutorials or poster set-up. Payment is given as a
check at the end of the meetings or when the student leaves Phoenix.

Once the program has been finalized, each registered bursary will be
contacted and allowed to choose the times and activities they would
like
to work. Many times they end up *working* sessions they would
attend anyway. There is an added bonus of $10 cash meal allotment for
each morning and/or afternoon session worked and a meeting shirt.

If anyone would like to participate in the bursary program, please
check the "I wish to apply for a student bursary" box on the
registration form. Bursary space is limited, so sign-up early.
Applicants for the bursaries must be members of MSA or MAS, enrolled
as
students at a recognized educational institution, and have paid the
registration fee.

For non-students, volunteers are also needed to help with
the above mentioned meeting activities. Although not paid on an
hourly
basis as the student bursaries, volunteers do receive the same cash
allotment for meals and a meeting shirt. Plus they have the
opportunity to interact more with the microscopy community as they
assist with meeting tasks.

If anyone has any questions about the bursary/volunteer program, or
would like to participate, please contact:

Amanda Lawrence
Institute for Imaging and Analytical Technologies
Mississippi State University
Box 9775
Mississippi State, MS 39762
662-325-7998
alawrence-at-i2at.msstate.edu

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Hi all.

Allow me to bring the following job opening (below) to your attention. Feel free forward the advertisement to any interested candidates who are not subscribing to this list. You can find further information about Camborne School of Mines at http://emps.exeter.ac.uk/csm/ and information about our analytical facilities at http://emps.exeter.ac.uk/csm/facilities/geochem-mineralogy/. It may also be of interest that the laboratories also have ICP-MS (since the webpages were updated).

Regards,

Jens

____________________________________________________

Dr. Jens C. Andersen
Senior lecturer
Camborne School of Mines
College of Engineering, Mathematics and Physical Sciences
University of Exeter
Cornwall Campus
Tremough
Penryn
TR10 9EZ
UNITED KINGDOM

Tel. +44 1326 371836
Fax. +44 1326 371859

http://www.exeter.ac.uk/csm

Visit the virtual Skaergaard intrusion: http://www.skaergaard.org

__________________________________________________________

Job title Experimental Officer & Technical Team Leader
Job reference R11073
Date posted 14/03/2012
Application closing date 15/04/2012
Location Cornwall
Salary �31,948 to �39,257
Package Generous holiday allowances, flexible working, pension scheme, car lease scheme and relocation package (if applicable).
Job category/type Admin, Professional, Managerial

Job description

College of Engineering, Mathematics and Physical Sciences
Camborne School of Mines, Penryn, Cornwall, United Kingdom

The post holder will be the technical team lead for the analytical labs at Camborne School of Mines (CSM) in support of research and teaching activities in the area of Responsible Mining, Sustainable Use of Natural Resources and Renewable Energy. The post holder will carry out research activities, support grant and income generation and provide teaching support to undergraduate students within the analytical lab in the area of electron-beam analysis.

Responsibilities include management of the Analytical laboratory/equipment area and staff; operation and maintenance of electron-beam analytical instruments including JEOL JXA 8200 electron microprobe and JEOL JSM 5600LV low-vacuum scanning electron microscope; working with academic staff on research; identifying grant and other income opportunities including contribution to grant applications and tender bids; tracking and managing research budget allocations; consultancy and collaboration with external partners; provision of design and technical advice for research projects and equipment used in electron microscopy and analysis; engaging with users to design and develop analytical protocols and experiments for use in research projects; plan, design, teach and supervise electron beam techniques and use of equipment to undergraduate and postgraduate students; providing undergraduate and postgraduate teaching support including delivering specified modules, marking, supporting student project work, providing demonstrations and workshops for student teaching.

For further information please contact Sam Livy, Assistant College Manager for Infrastructure and Technical Services, e-mail S.A.Livy-at-exeter.ac.uk or telephone (01392) 724477.

Priority consideration will generally be given to internal applicants.

To view the Job Description and Person Specification document please follow this link http://admin.exeter.ac.uk/personnel/jobs/R11073.pdf

The University of Exeter is an equal opportunity employer which is 'Positive About Disabled People': if you have a disability, you should mention this in your application. Whilst all applicants will be judged on merit alone, we particularly welcome applications from groups currently underrepresented in the workforce.

==============================Original Headers==============================
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21, 28 -- Subject: EPMA - Job opportunity for electron microprobe operator/lab manager
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A user would like to make his coated grids less hydrophobic and more
hydrophilic. These are 3.0mm copper grids coated with formvar and carbon

Any ideas?

Fred Hayes
Manager
Interdisciplinary Center for Electron Microscopy (ICEM) / Kemper Hall
Facility, room 0108
Det of Chemical Engineering and Material Sciences
3118 Bainer Hall
Univ of CA Davis
Davis CA 95616
530-752-0284 office

==============================Original Headers==============================
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8, 23 -- To: {Microscopy-at-microscopy.com}
8, 23 -- Cc: "Feng Jiang" {fjiang-at-ucdavis.edu}
8, 23 -- Subject: making coated grids less hydrophobic
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Email: fsmsc-at-hotmail.com Name: Fanny Chu

Organization: James Hogg Research Centre/Heart & Lung Institute

Title-Subject: [Filtered] LaB6 operating gun vacuum limit

Message: Can anyone please share your experience of using the LaB6 filament on your TEM,
particularly if you have an FEI Tecnai 12? What operating vacuum in real life is required so you
don't blow the LaB6?
Background:
- My old LaB6 (15 micron tip microflat) was purchased from Barry Scientific (apparently no longer in
service). In spite of maintenance shut off, building power shut off and vacuum crashes during
specimen insertion, it lasted 5.5 years with an operating (gun)vacuum of 6-13(log scale, or { {10e-7
torr);

- The newly replaced LaB6 (bought from Barry Scientific and stored since 2008) only lasted 4 months
and blew not when the scope was busiest. BSc reassured me that their filaments can be stored for
long time;

- My Tecnai 12 TEM had its column opened twice for O rings replacement in the last several weeks.
One more set of O rings to replace since vacuum is still sub-optimal;

- Vendor was convinced that a log value under 30 should be fine and my scope was operating between
19-25. When the LaB6 blew, the operating vacuum was at 20. BTW, a log scale 22 is equivalent 0.35e-6;
- The Electron Microscopy Sciences document indicates that for LaB6, operation vacuum should be at
10e-7, work function (eV) should be ~2.70. My operating eV was ~2.30.

Is operating at a vacuum between 20-30 pushing the limit? Any suggestion, even about handling,
maintenance, storage, etc would be appreciated.

Sincerely,

Fanny Chu
Ultrastructural Imaging,
UBC James Hogg Research Centre/
Institute of Heart and Lung,
St. Paul's Hospital,
Rm 166, 1081 Burrard St, Vancouver, BC, Canada
604) 682-2344, x 62712

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I've done this. No issues other than the usual getting-the-best-image issues.
Phil

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}
} Email: cmeyer911-at-yahoo.com Name: Chris Meyer
}
} Title-Subject: [Filtered] Immersion lens imaging of nanotubes
}
} Message: Are there any issues imaging carbon nanotubes in an SEM
} under an immersion lens setting?
} Thanks.
}

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

==============================Original Headers==============================
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Fred,
My colleague Chris Woodcock at UMass Amherst makes his carbon
coated grids more hydrophillic by exposing them to a plasma for 30
sec or so (I am not sure of the exact timing). Our microscopy core
has a plasma cleaner that is ideal for this purpose. I use this
frequently to make slides and coverslips hydrophillic but it works
for grids too. Put the grids or whatever in the small chamber, it is
evacuated modestly (I think to 1%? maybe 0.1%, anyway just with a
simple pump), and then the plasma is struck. About 1 min later turn
off and you have nice hydrophillic samples. I believe this works by
activated radicals in the plasma attacking the hydocarbon film that
coats every surface in the universe.

Hope this helps,
Tobias
}
}
} A user would like to make his coated grids less hydrophobic and more
} hydrophilic. These are 3.0mm copper grids coated with formvar and carbon
}
} Any ideas?
}
}
}
} Fred Hayes
} Manager
} Interdisciplinary Center for Electron Microscopy (ICEM) / Kemper Hall
} Facility, room 0108
} Det of Chemical Engineering and Material Sciences
} 3118 Bainer Hall
} Univ of CA Davis
} Davis CA 95616
} 530-752-0284 office
}

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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Email: dservando86-at-yahoo.com Name: Donya Servando

Organization: San Joaquin Delta College EM Dept.

Title-Subject: [Filtered] TEM sample prep using freezing methods

Message: I am a student at Delta College in the EM program and I am currently working on a special
project that involves subjecting my sample (duck liver) to liquid nitrogen for freeze drying and
then processing the sample for TEM analysis.

My problem is that our school is not equipped with instrumentation to do cryo work, let alone pretty
basic methods for freeze drying. The only tools I have is LN2, a brass bucket to "plunge" my
samples into, and a Varian Vacuum Evaporater. My question is, after the sample has "thawed" in the
VE after a few days, what is the next best step for TEM prep? Do I go straight into infiltration
with 100% acetone and then into the ratios of resin:acetone? If the sample has not been put into
any heavy metal salt such as UA or OsO4, what about contrast issues? Are there any steps I should
take BEFORE plunging my sample into LN2 and then left to thaw under vacuum?

So far, the closest I have come to finding some form of a standard protocol for freeze drying is by
molecular distillation and Kent McDonald's method for HPF/QFS. However, MDD's are no longer
manufactured (we don't have one anyway) and we do not have a high pressure freezer for the HPF/QFS.
Does anyone have any "innovative" ideas that I could try out? Basically, I need to freeze dry a
sample (with or without some form contrast element; whichever you all would think is a better
choice) and the embed it into resin for TEM. No special tricks or tools, just the basics! I would
appreciate any advice since as of right now, I am pretty much taking bits and pieces from different
protocols and "winging it".

Thanks for any help,
Donya

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Dear Chris,

We never had problems with nanotubes using with in-lens for imaging. Just once, our user was analyzing some kind of loose fiber and a small fiber got inside the lens and start causing problems with the imaging. But this fiber was enormous in size if compared with nanotubes. After that we start to ask our users to use an air blower to remove any loose piece of sample on the carbon tape.

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil
________________________________________
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Email: cmeyer911-at-yahoo.com Name: Chris Meyer

Title-Subject: [Filtered] Immersion lens imaging of nanotubes

Message: Are there any issues imaging carbon nanotubes in an SEM under an immersion lens setting?
Thanks.

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Fanny,
To me (and I imagine to many other non-Tecnai people) the log reading is
pretty meaningless. However, if a log scale of 22 equals 0.35e-6T, that
would be 3.5e-7T which is OK. Not terrific, but OK. If you meant 3.5e-6,
that is terrible and could explain the short filament life.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
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Email: fsmsc-at-hotmail.com Name: Fanny Chu

Organization: James Hogg Research Centre/Heart & Lung Institute

Title-Subject: [Filtered] LaB6 operating gun vacuum limit

Message: Can anyone please share your experience of using the LaB6 filament
on your TEM,
particularly if you have an FEI Tecnai 12? What operating vacuum in real
life is required so you
don't blow the LaB6?
Background:
- My old LaB6 (15 micron tip microflat) was purchased from Barry Scientific
(apparently no longer in
service). In spite of maintenance shut off, building power shut off and
vacuum crashes during
specimen insertion, it lasted 5.5 years with an operating (gun)vacuum of
6-13(log scale, or { {10e-7
torr);

- The newly replaced LaB6 (bought from Barry Scientific and stored since
2008) only lasted 4 months
and blew not when the scope was busiest. BSc reassured me that their
filaments can be stored for
long time;

- My Tecnai 12 TEM had its column opened twice for O rings replacement in
the last several weeks.
One more set of O rings to replace since vacuum is still sub-optimal;

- Vendor was convinced that a log value under 30 should be fine and my scope
was operating between
19-25. When the LaB6 blew, the operating vacuum was at 20. BTW, a log scale
22 is equivalent 0.35e-6;
- The Electron Microscopy Sciences document indicates that for LaB6,
operation vacuum should be at
10e-7, work function (eV) should be ~2.70. My operating eV was ~2.30.

Is operating at a vacuum between 20-30 pushing the limit? Any suggestion,
even about handling,
maintenance, storage, etc would be appreciated.

Sincerely,

Fanny Chu
Ultrastructural Imaging,
UBC James Hogg Research Centre/
Institute of Heart and Lung,
St. Paul's Hospital,
Rm 166, 1081 Burrard St, Vancouver, BC, Canada
604) 682-2344, x 62712

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On Mar 15, 2012, at 3:47 PM, fahayes-at-ucdavis.edu wrote:

} A user would like to make his coated grids less hydrophobic and more
} hydrophilic. These are 3.0mm copper grids coated with formvar and
} carbon
}
} Any ideas?

Dear Fred,
Glow-discharging the grid for a minute, give or take, is one process,
and coating the grid with poly-L-lysine is another.
Yours,
Bill

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If your tissue fixation method is plunging a chunk of liver into liquid nitrogen, you don't need to worry about how to freeze-dry it because the preservation will be so bad, it isn't worth looking at. Liquid nitrogen might seem like a fast way to freeze something but it isn't fast enough - vaporization of the nitrogen forms an insulating layer that slows freezing to milliseconds. This results in ice crystals that are big enough to disrupt tissue structure. You need to use high pressure freezing with a really expensive instrument or a metal mirror device that is moderately expensive. If you have well frozen tissue, freeze-substitution is much easier and more common than freeze-drying. For most purposes it is satisfactory. Freeze-substitution can be done with homemade contraptions involving ice chests and dry ice.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Email: dservando86-at-yahoo.com Name: Donya Servando

Organization: San Joaquin Delta College EM Dept.

Title-Subject: [Filtered] TEM sample prep using freezing methods

Message: I am a student at Delta College in the EM program and I am currently working on a special project that involves subjecting my sample (duck liver) to liquid nitrogen for freeze drying and then processing the sample for TEM analysis.

My problem is that our school is not equipped with instrumentation to do cryo work, let alone pretty basic methods for freeze drying. The only tools I have is LN2, a brass bucket to "plunge" my samples into, and a Varian Vacuum Evaporater. My question is, after the sample has "thawed" in the VE after a few days, what is the next best step for TEM prep? Do I go straight into infiltration with 100% acetone and then into the ratios of resin:acetone? If the sample has not been put into any heavy metal salt such as UA or OsO4, what about contrast issues? Are there any steps I should take BEFORE plunging my sample into LN2 and then left to thaw under vacuum?

So far, the closest I have come to finding some form of a standard protocol for freeze drying is by molecular distillation and Kent McDonald's method for HPF/QFS. However, MDD's are no longer manufactured (we don't have one anyway) and we do not have a high pressure freezer for the HPF/QFS.
Does anyone have any "innovative" ideas that I could try out? Basically, I need to freeze dry a sample (with or without some form contrast element; whichever you all would think is a better
choice) and the embed it into resin for TEM. No special tricks or tools, just the basics! I would appreciate any advice since as of right now, I am pretty much taking bits and pieces from different protocols and "winging it".

Thanks for any help,
Donya

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Donya,

You might want to try the freezing methods described in these publications:

Self-pressurized rapid freezing (SPRF): a novel cryofixation method for specimen preparation in electron microscopy by J.L.M. LEUNISSEN & H. YI at: www.aurion.nl/pdf/LeunissenYi2009.pdf

Freezing in sealed capillaries for preparation of frozen hydrated sections by S. YAKOVLEV and K.H. DOWNING Journal of Microscopy Volume 244, Issue 3, Article first published online: 3 OCT 2011 at http://onlinelibrary.wiley.com/doi/10.1111/j.1365-2818.2011.03530.x/pdf

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

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} Title-Subject: [Filtered] TEM sample prep using freezing methods
}
} Message: I am a student at Delta College in the EM program and I am currently working on a special
} project that involves subjecting my sample (duck liver) to liquid nitrogen for freeze drying and
} then processing the sample for TEM analysis.
}
} My problem is that our school is not equipped with instrumentation to do cryo work, let alone pretty
} basic methods for freeze drying. The only tools I have is LN2, a brass bucket to "plunge" my
} samples into, and a Varian Vacuum Evaporater. My question is, after the sample has "thawed" in the
} VE after a few days, what is the next best step for TEM prep? Do I go straight into infiltration
} with 100% acetone and then into the ratios of resin:acetone? If the sample has not been put into
} any heavy metal salt such as UA or OsO4, what about contrast issues? Are there any steps I should
} take BEFORE plunging my sample into LN2 and then left to thaw under vacuum?
}
} So far, the closest I have come to finding some form of a standard protocol for freeze drying is by
} molecular distillation and Kent McDonald's method for HPF/QFS. However, MDD's are no longer
} manufactured (we don't have one anyway) and we do not have a high pressure freezer for the HPF/QFS.
} Does anyone have any "innovative" ideas that I could try out? Basically, I need to freeze dry a
} sample (with or without some form contrast element; whichever you all would think is a better
} choice) and the embed it into resin for TEM. No special tricks or tools, just the basics! I would
} appreciate any advice since as of right now, I am pretty much taking bits and pieces from different
} protocols and "winging it".
}
} Thanks for any help,
} Donya
}
} Login Host: 99.63.218.76

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Donya

there are ways to improve the initial freezing rate of liquid nitrogen but you will need to exercise care because anything that improves the freezing rate of samples does the same to you and can cause burns to skin which a simple splash wit drops of liquid nitrogen wouldn't normally do.

The simplest way to improve cooling rates of liquid nitrogen is to "slush" it. This is done by pumping it down in a partial vacuum until until it starts to form liquid nitrogen slush (solid nitrogen) by latent heat of evaporation. The process often needs to be repeated two or three times quickly to supercool the nitrogen properly. However (and this is a big however) I would never leave an untrained graduate project student alone to do this - I would always closely supervise because of the hazards of the violent boiling of nitrogen and its supercooling properties. Other methods involve various gases which can be liquified by nitrogen. the sample is then plunged into the liquified gas rather than the nitrogen. Freon used to be used but is now banned and most of the rest are highly flammable and would need careful handling a fume hood.

Another simple way of improving the quality of freezing is to use a cryoprotectant such as sucrose, glycerol or PVP. These are best done after fixation though so may be of limited use if you are trying to completely immobilize cell contents for x-ray micro-analysis.

You may need to do a bit more reading around to decide which is the best route and be particulalrly wary of some of the hazards of the cryogens. I came across this while googling em cryoprotectants:
http://books.google.co.uk/books?id=dqP8FCEiMGgC&pg=PA205&lpg=PA205&dq=em+cryoprotectants&source=bl&ots=C_Mr5M9K7N&sig=d3zLR5k-F3DcCbJ8FA7d4V8YqUU&hl=en&sa=X&ei=EGljT7uqHoK90QW_vqS_CA&ved=0CD0Q6AEwBA#v=onepage&q=em%20cryoprotectants&f=false

It gives a good summary even if it may not help.

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: PhillipsT-at-missouri.edu [PhillipsT-at-missouri.edu]
Sent: 16 March 2012 15:45
To: Malcolm Haswell

If your tissue fixation method is plunging a chunk of liver into liquid nitrogen, you don't need to worry about how to freeze-dry it because the preservation will be so bad, it isn't worth looking at. Liquid nitrogen might seem like a fast way to freeze something but it isn't fast enough - vaporization of the nitrogen forms an insulating layer that slows freezing to milliseconds. This results in ice crystals that are big enough to disrupt tissue structure. You need to use high pressure freezing with a really expensive instrument or a metal mirror device that is moderately expensive. If you have well frozen tissue, freeze-substitution is much easier and more common than freeze-drying. For most purposes it is satisfactory. Freeze-substitution can be done with homemade contraptions involving ice chests and dry ice.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Email: dservando86-at-yahoo.com Name: Donya Servando

Organization: San Joaquin Delta College EM Dept.

Title-Subject: [Filtered] TEM sample prep using freezing methods

Message: I am a student at Delta College in the EM program and I am currently working on a special project that involves subjecting my sample (duck liver) to liquid nitrogen for freeze drying and then processing the sample for TEM analysis.

My problem is that our school is not equipped with instrumentation to do cryo work, let alone pretty basic methods for freeze drying. The only tools I have is LN2, a brass bucket to "plunge" my samples into, and a Varian Vacuum Evaporater. My question is, after the sample has "thawed" in the VE after a few days, what is the next best step for TEM prep? Do I go straight into infiltration with 100% acetone and then into the ratios of resin:acetone? If the sample has not been put into any heavy metal salt such as UA or OsO4, what about contrast issues? Are there any steps I should take BEFORE plunging my sample into LN2 and then left to thaw under vacuum?

So far, the closest I have come to finding some form of a standard protocol for freeze drying is by molecular distillation and Kent McDonald's method for HPF/QFS. However, MDD's are no longer manufactured (we don't have one anyway) and we do not have a high pressure freezer for the HPF/QFS.
Does anyone have any "innovative" ideas that I could try out? Basically, I need to freeze dry a sample (with or without some form contrast element; whichever you all would think is a better
choice) and the embed it into resin for TEM. No special tricks or tools, just the basics! I would appreciate any advice since as of right now, I am pretty much taking bits and pieces from different protocols and "winging it".

Thanks for any help,
Donya

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The capillary method is impressive for yeast or C. elegans or bacteria but not practical for liver. One would need a more complicated biopsy device with a thin lumen and need to work fast to avoid structural changes from the biopsy step. The Yakolev and Dowling paper used cryoprotectants which are more problematic with a tissue slice. Quick freezing is the best fixation method but it is such a pain in the neck, not many people really do it. The number of papers per year employing it is not overwhelming. I say this as someone who co-developed a metal mirror device (sold under the name "Gentleman Jim" by my brilliant thesis adviser Alan Boyne) and someone who bought one of Leica's High Pressure Freezing instruments (based on a great design by Jan Studer).

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: rcsencsits-at-lbl.gov [mailto:rcsencsits-at-lbl.gov]
Sent: Friday, March 16, 2012 11:07 AM
To: Phillips, Thomas E.

Donya,

You might want to try the freezing methods described in these publications:

Self-pressurized rapid freezing (SPRF): a novel cryofixation method for specimen preparation in electron microscopy by J.L.M. LEUNISSEN & H. YI at: www.aurion.nl/pdf/LeunissenYi2009.pdf

Freezing in sealed capillaries for preparation of frozen hydrated sections by S. YAKOVLEV and K.H. DOWNING Journal of Microscopy Volume 244, Issue 3, Article first published online: 3 OCT 2011 at http://onlinelibrary.wiley.com/doi/10.1111/j.1365-2818.2011.03530.x/pdf

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

On Mar 16, 2012, at 6:08 AM, microscopylistserver-noreply-at-microscopy.com wrote:

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} Email: dservando86-at-yahoo.com Name: Donya Servando
}
} Organization: San Joaquin Delta College EM Dept.
}
} Title-Subject: [Filtered] TEM sample prep using freezing methods
}
} Message: I am a student at Delta College in the EM program and I am
} currently working on a special project that involves subjecting my
} sample (duck liver) to liquid nitrogen for freeze drying and then processing the sample for TEM analysis.
}
} My problem is that our school is not equipped with instrumentation to
} do cryo work, let alone pretty basic methods for freeze drying. The
} only tools I have is LN2, a brass bucket to "plunge" my samples into,
} and a Varian Vacuum Evaporater. My question is, after the sample has
} "thawed" in the VE after a few days, what is the next best step for
} TEM prep? Do I go straight into infiltration with 100% acetone and
} then into the ratios of resin:acetone? If the sample has not been put into any heavy metal salt such as UA or OsO4, what about contrast issues? Are there any steps I should take BEFORE plunging my sample into LN2 and then left to thaw under vacuum?
}
} So far, the closest I have come to finding some form of a standard
} protocol for freeze drying is by molecular distillation and Kent
} McDonald's method for HPF/QFS. However, MDD's are no longer manufactured (we don't have one anyway) and we do not have a high pressure freezer for the HPF/QFS.
} Does anyone have any "innovative" ideas that I could try out?
} Basically, I need to freeze dry a sample (with or without some form
} contrast element; whichever you all would think is a better
} choice) and the embed it into resin for TEM. No special tricks or
} tools, just the basics! I would appreciate any advice since as of
} right now, I am pretty much taking bits and pieces from different protocols and "winging it".
}
} Thanks for any help,
} Donya
}
} Login Host: 99.63.218.76

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Email: fsmsc-at-hotmail.com Name: Fanny Chu

Organization: James Hogg Research Centre/Heart & Lung Institute

Title-Subject: [Filtered] Mystery solved - re: LaB6 operating gun vacuum limit

Message: The mystery about my LaB6 was solved.

It's embarrassing. I have just found out this moment that the filament that blew was a tungsten one,
not a LaB6!!

Somehow there was mis-communication so a tungsten one was used.

Many thanks to Ken Converse for his input.

Fanny Chu
Ultrastructural Imaging,
UBC James Hogg Research Centre/
Institute of Heart and Lung,
St. Paul's Hospital,
Rm 166, 1081 Burrard St, Vancouver, BC, Canada
(604) 682-2344, x 62712

Fanny, we have two FEI CM20 TEMs that use LaB6 cathodes. We have used
both Denka and Kimball Physics cathodes and get very long life ( many
years).
The key is to be sure the vacuum on the IGP is good (we never turn up
the cathode when IGP is } 20.) Even more important is to saturate
slowly (I wrote a plug-in for DigitalMicrograph that turns it up in 10
sec increments) and to check the saturation after 15 min - the cathode
continues to heat and can oversaturate. On our cryoscope, we have a
turbo-pumped air lock and see no major jump in IGP on sample exchange,
so I will change a sample with the cathode backed off a few clicks;
the other scope only uses the mechanical pump and we get frequent
vacuum jumps. I turn down the cathode and wait a bit before sample
exchange. I'd rather take a few moments to have a sip of coffee while
I wait than to have to spend the time changing the cathode and
conditioning our gun to 200 kV...

Best regards,
John

} Email: fsmsc-at-hotmail.com Name: Fanny Chu
} Organization: James Hogg Research Centre/Heart & Lung Institute
} Title-Subject: [Filtered] LaB6 operating gun vacuum limit
}
} Message: Can anyone please share your experience of using the LaB6 filament on your TEM,
} particularly if you have an FEI Tecnai 12? What operating vacuum in real life is required so you
} don't blow the LaB6?
} Background:
} - My old LaB6 (15 micron tip microflat) was purchased from Barry Scientific (apparently no longer in
} service). In spite of maintenance shut off, building power shut off and vacuum crashes during
} specimen insertion, it lasted 5.5 years with an operating (gun)vacuum of 6-13(log scale, or { {10e-7
} torr);
}
} - The newly replaced LaB6 (bought from Barry Scientific and stored since 2008) only lasted 4 months
} and blew not when the scope was busiest. BSc reassured me that their filaments can be stored for
} long time;
}
} - My Tecnai 12 TEM had its column opened twice for O rings replacement in the last several weeks.
} One more set of O rings to replace since vacuum is still sub-optimal;
}
} - Vendor was convinced that a log value under 30 should be fine and my scope was operating between
} 19-25. When the LaB6 blew, the operating vacuum was at 20. BTW, a log scale 22 is equivalent 0.35e-6;
} - The Electron Microscopy Sciences document indicates that for LaB6, operation vacuum should be at
} 10e-7, work function (eV) should be ~2.70. My operating eV was ~2.30.
}
} Is operating at a vacuum between 20-30 pushing the limit? Any suggestion, even about handling,
} maintenance, storage, etc would be appreciated.
}
} Sincerely,
}
} Fanny Chu
} Ultrastructural Imaging,
} UBC James Hogg Research Centre/
} Institute of Heart and Lung,
} St. Paul's Hospital,
} Rm 166, 1081 Burrard St, Vancouver, BC, Canada
} 604) 682-2344, x 62712

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Hi Donya,

Why do you want to do the freezing? Do you plan on doing some sort of gold immuno-labeling? Just
wondering why you are trying to do the freezing when you don't really have the equipment. Your
sample is not going to have good morphology - you are going to have all sorts of freezing damage.

Kind Regards,
Peggy Bisher

Applications Specialist
Gatan, Inc
780 Commonwealth Drive
Warrendale, PA 15086 USA
(724) 779-2577 Direct
(724) 234-9207 Cell

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-----Original Message-----
} From: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Friday, March 16, 2012 9:09 AM
To: Margaret Bisher

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Email: dservando86-at-yahoo.com Name: Donya Servando

Organization: San Joaquin Delta College EM Dept.

Title-Subject: [Filtered] TEM sample prep using freezing methods

Message: I am a student at Delta College in the EM program and I am currently working on a special
project that involves subjecting my sample (duck liver) to liquid nitrogen for freeze drying and
then processing the sample for TEM analysis.

My problem is that our school is not equipped with instrumentation to do cryo work, let alone pretty
basic methods for freeze drying. The only tools I have is LN2, a brass bucket to "plunge" my
samples into, and a Varian Vacuum Evaporater. My question is, after the sample has "thawed" in the
VE after a few days, what is the next best step for TEM prep? Do I go straight into infiltration
with 100% acetone and then into the ratios of resin:acetone? If the sample has not been put into
any heavy metal salt such as UA or OsO4, what about contrast issues? Are there any steps I should
take BEFORE plunging my sample into LN2 and then left to thaw under vacuum?

So far, the closest I have come to finding some form of a standard protocol for freeze drying is by
molecular distillation and Kent McDonald's method for HPF/QFS. However, MDD's are no longer
manufactured (we don't have one anyway) and we do not have a high pressure freezer for the HPF/QFS.
Does anyone have any "innovative" ideas that I could try out? Basically, I need to freeze dry a
sample (with or without some form contrast element; whichever you all would think is a better
choice) and the embed it into resin for TEM. No special tricks or tools, just the basics! I would
appreciate any advice since as of right now, I am pretty much taking bits and pieces from different
protocols and "winging it".

Thanks for any help,
Donya

Login Host: 99.63.218.76
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Donya,
Look into small diameter copper tube freezing, sealed at both ends after you get the tissue into it.
Apparently the small volume can mimick the high pressure of an HPF machine when you plunge into
liquid nitrogen. If you dont have the high pressure, when you freeze samples the water will cause
ice crystal damage.

good luck,

Michael Delannoy

----- Original Message -----
} From: microscopylistserver-noreply-at-microscopy.com

Hi

I observed much NTs grown on different kind of substrates and had no
problemes.
But a collegue observes other kind of growings, whith Ni or Fe as
catalysator and where the NTs are free to move, and we must regulary
clean the OL bore from the dust which is traped by the magnetic field.
I require since to pass the sample holder under a strong magnet (from a
old hard drive), to catch all what could be free to fly away. It's a
little bit better, but not perfect.

Hope it helps

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Mat�riaux de Strasbourg
D�partement Surfaces et Interfaces
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail : Jacques.Faerber-at-ipcms.unistra.fr

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} Email: cmeyer911-at-yahoo.com Name: Chris Meyer
}
} Title-Subject: [Filtered] Immersion lens imaging of nanotubes
}
} Message: Are there any issues imaging carbon nanotubes in an SEM under an immersion lens setting?
} Thanks.
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Email: fanny.chu-at-hli.ubc.ca Name: Fanny Chu

Organization: James Hogg Research Centre / Heart & Lung Institute

Title-Subject: [Filtered] LaB6 operating gun vacuum limit

Message: Thank you John for your very helpful input.

What you said about keeping IGP vacuum at {20 and checking saturation is so true. At least that was
my own experience of having the old LaB6 that lasted this long.

Best regards,
Fanny

Fanny, we have two FEI CM20 TEMs that use LaB6 cathodes. We have used
both Denka and Kimball Physics cathodes and get very long life ( many
years).
The key is to be sure the vacuum on the IGP is good (we never turn up
the cathode when IGP is } 20.) Even more important is to saturate
slowly (I wrote a plug-in for DigitalMicrograph that turns it up in 10
sec increments) and to check the saturation after 15 min - the cathode
continues to heat and can oversaturate. On our cryoscope, we have a
turbo-pumped air lock and see no major jump in IGP on sample exchange,
so I will change a sample with the cathode backed off a few clicks;
the other scope only uses the mechanical pump and we get frequent
vacuum jumps. I turn down the cathode and wait a bit before sample
exchange. I'd rather take a few moments to have a sip of coffee while
I wait than to have to spend the time changing the cathode and
conditioning our gun to 200 kV...
Best regards,
John
} Email: fsmsc-at-hotmail.com Name: Fanny Chu
} Organization: James Hogg Research Centre/Heart & Lung Institute
} Title-Subject: [Filtered] LaB6 operating gun vacuum limit
}
} Message: Can anyone please share your experience of using the LaB6 filament on your TEM,
} particularly if you have an FEI Tecnai 12? What operating vacuum in real life is required so you
} don't blow the LaB6?
} Background:
} - My old LaB6 (15 micron tip microflat) was purchased from Barry Scientific (apparently no longer in
} service). In spite of maintenance shut off, building power shut off and vacuum crashes during
} specimen insertion, it lasted 5.5 years with an operating (gun)vacuum of 6-13(log scale, or { {10e-7
} torr);
}
} - The newly replaced LaB6 (bought from Barry Scientific and stored since 2008) only lasted 4 months
} and blew not when the scope was busiest. BSc reassured me that their filaments can be stored for
} long time;
}
} - My Tecnai 12 TEM had its column opened twice for O rings replacement in the last several weeks.
} One more set of O rings to replace since vacuum is still sub-optimal;
}
} - Vendor was convinced that a log value under 30 should be fine and my scope was operating between
} 19-25. When the LaB6 blew, the operating vacuum was at 20. BTW, a log scale 22 is equivalent 0.35e-6;
} - The Electron Microscopy Sciences document indicates that for LaB6, operation vacuum should be at
} 10e-7, work function (eV) should be ~2.70. My operating eV was ~2.30.
}
} Is operating at a vacuum between 20-30 pushing the limit? Any suggestion, even about handling,
} maintenance, storage, etc would be appreciated.
}
} Sincerely,
}
} Fanny Chu
} Ultrastructural Imaging,
} UBC James Hogg Research Centre/
} Institute of Heart and Lung,
} St. Paul's Hospital,
} Rm 166, 1081 Burrard St, Vancouver, BC, Canada
} 604) 682-2344, x 62712

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Hi Ben,
We have a UC6, also a few years old, also showed such problems. Apparently
the component parts have to be cleaned and new lubricant applied on a
fairly regular basis. So I was told by the engineer who fixed our one. It
wasn't cheap and we were told we should have had it serviced every year,
but we aren't doing that - too expensive. You may need to invest in a
full service for each.
cheers,
Rosemary

Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
E rosemary.white-at-csiro.au

On 20/03/12 9:24 AM, "microscopylistserver-noreply-at-microscopy.com"
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Position: TEM Facility Manager in the Materials Science & Engineering department at North Carolina State University, Raleigh NC

Job Description:

The position will have primary responsibility for management and operation of the Materials Science & Engineering Transmission Electron Microscopy Facility at NCSU, which houses a JEOL 2010F S/TEM, JEOL 2000FX LaB6, JEOL Touchscope SEM and auxiliary equipment, including energy dispersive x-ray spectrometers, electron energy loss spectrometer, CCD cameras and an extensive sample preparation laboratory.

Duties and Responsibilities:

� Oversee training of students, faculty, and external researchers on SEM and TEM-related equipment and software � establish facility policies for user training and oversight; train users to operate the TEMs and auxiliary equipment with the ultimate goal of teaching users to perform basic microscope functions safely and independently; assist users with performing advanced microscopy techniques;
� Interact with external users to provide microscope training and/or materials analysis via a research service agreement
� Ensure that instruments are properly functioning and calibrated at all times and as necessary; coordinate service by self, MSE support staff and/or the product vendors. This includes establishing routine maintenance and calibration schedules for the microscopes and all detectors.
� Schedule microscope usage and establish and enforce user policies that maintain an orderly, clean and safe laboratory.
� Assist in the design and implementation of user seminars or short courses. These activities will be designed to inform users of recent advancements in electron microscopy techniques and data quantification with the aim of improving the quality of graduate education and research at the University.
� Stay abreast of software and hardware developments by reading current literature and attending relevant technical conferences and short courses
� Administrate and maintain the acquisition computers and coordinate routine back ups, virus checks, etc. Some computer support is available from MSE. Keep users informed of the analysis programs available.
� Manage other TEM staff, who will assist with daily upkeep of the facility and user training.
� Communicate the need for instrument upgrades and develop strategies and proposals for acquiring equipment upgrades or replacements.
� Work with the University Wide TEM Steering Committee

Minimum Education/Experience:

a) MS in Materials, Physical Sciences or related engineering field and a strong background in the theory and practice of TEM microcharacterizaton AND minimum of three years post-graduate experience

OR

b) Ph.D in Materials, Physical Sciences or related engineering field.

Departmental Required Skills :

Experience with vacuum technology, electronics, and computers. Strong interpersonal and communications skills are essential.

To apply or for more information see:

https://jobs.ncsu.edu/postings/5362

AA/EOE. In addition, NC State welcomes all persons without regard to sexual orientation. Persons with disabilities requiring accommodations in the application and interview process please call (919) 515-3148. Final candidates are subject to criminal & sex offender background checks. Some vacancies also require credit or motor vehicle checks. If highest degree is from an institution outside of the U.S., final candidates are required to have their degree verified at www.wes.org. Degree must be obtained prior to start date. NC State University participates in E-Verify. Federal law requires all employers to verify the identity and employment eligibility of all persons hired to work in the United States.

James LeBeau
Assistant Professor
Department of Materials Science & Engineering
North Carolina State University
tel: (919) 515.5049
jmlebeau at ncsu.edu

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18, 33 -- Subject: Job Posting: Electron Microscopy Facility Manager
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Dear All:

I am in need of a gold conjugated anti-FLAG antibody. Does anyone know a
source to purchase from? Thank you all in advance.

Hong

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Vickie:

1) If you use glycerin as a cryoprotectant you will inhibit etching.
2) Plunging into LN2 is not recommended for direct freezing your samples.
Use Propane or Ethane as a cryogen.
3) You cannot control the fracture plane.

All this is information is well published. Try a literature search: Dr. Joy
Frank, UCLA. She and her tech, Tony Mottino published many papers on FF/DE
of Cardiac Muscle using simple freezing methods.

Al Coritz
Applications & Service Manager
EMS Technical Service
www.emsdiasum.com

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Email: vakimler-at-med.wayne.edu Name: Vickie Kimler

Organization: Wayne State University School of Medicine

Title-Subject: [Filtered] Freeze-Fracture Deep-Etch SEM

Message: I have cardiac muscle tissue (~1-2 mm2) chunks that I have fixed in
glutaraldehyde and paraformaldehyde at 4 degrees C for about six hours on
Monday. The samples were then placed into phosphate buffered saline at 4
degrees C and are nutating.

I plan to prep for freeze-fracture deep-etch SEM on Friday. Would it be a
good idea to add 30% glycerol to buffer before plunging it in LN2? Any other
suggestions to reduce ice crystallization, enhance sharp fracture planes,
perform effective etching, etc.? We are using an Alto 2500 cryotransfer
device.

Thank you,
Vickie

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Does anyone have a manual for the Denton DESK II sputter coater?

Thank you

Fred Hayes
Manager
Interdisciplinary Center for Electron Microscopy (ICEM) / Kemper Hall
Facility, room 0108
Det of Chemical Engineering and Material Sciences
3118 Bainer Hall
Univ of CA Davis
Davis CA 95616
530-752-0284 office

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Fred,

I have one I could photocopy, maybe scan and send as a pdf.

Phil

} Does anyone have a manual for the Denton DESK II sputter coater?
}
} Thank you
}
} Fred Hayes
} Manager
} Interdisciplinary Center for Electron Microscopy (ICEM) / Kemper Hall
} Facility, room 0108
} Det of Chemical Engineering and Material Sciences
} 3118 Bainer Hall
} Univ of CA Davis
} Davis CA 95616
} 530-752-0284 office

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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http://seankampshoff.com/images/truesdal.htm

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The Company:
Our client is a leading developer and producer of innovative high-tech
precision optics systems for the analysis of microstructures.� As one of
the market leaders in each of the fields of Microscopy, Confocal Laser
Scanning Microscopy, Imaging Systems, Specimen Preparation and Medical
Equipment.� Comprising nine manufacturing facilities in seven countries,
sales and service companies in 20 countries and an international network
of dealers, the company is represented in over 100 countries.

The Opportunity:
The company currently has an opening for a Research Microscopy Specialist
to be based in Irvine, California .� All applicants must not be adverse to
travel, as this is a position that may require you to travel when
necessary.

Base: $50-70k per year + bonus/commission

Other: Full benefits - 401k program/matching - Car allowance - Company
cell phone - Laptop computer - Home office set-up - Expenses

Primary Responsibilities:
As a key member of the sales team, the purpose of this position is to
promote sales of the company's Widefield & Advanced Fluorescence product
line in the assigned region.� The primary function is to execute domestic
sales plans, recommend strategies to improve the sale of Widefield &
Advanced Fluorescence products in the territory and aid in the
implementation of these strategies in line with Marketing, Corporate
growth, profitability and mission objectives.

Additional Responsibilities:
- Schedule product seminars or training sessions in concert with company
Product Management in the assigned region
- Participate in all activities that will enhance the awareness,
acceptance and sale of confocal products
- Conduct analysis and performance evaluation of all sales by use of the
company's Customer Resource Management tool
- Work with Sales Manager to develop strategies designed to increase sales
of confocal products; Specifically, the RMS will recommend product,
pricing, advertising and sales promotion strategies
- Coordinates the generation of new product ideas or modifications,
performs analysis and preliminary screening of these ideas and recommends
the appropriate ones to the Area Sales Manager for acceptance or rejection

Education and Experience Required:
BA/BS or MS degree in physics or chemistry or a related scientific field
is highly preferred.� Experience working with microscopy products is
required and a minimum of 2 years related microscopy experience in a
laboratory is essential.� In-depth knowledge of the microscopy market is
also highly desirable.� Sales experience in selling products of a
technical nature is desirable but not mandatory.� The individual must be
highly numerate as pricing, quoting and reviewing commercial terms will be
a daily routine. Strong computer skills and experience with CRM systems
(Maximizer, ACT, Goldmine, Siebel, etc).� Strong technical,
electrochemical, optical and mechanical aptitude.

If you meet and/or exceed the experience criteria, please submit your
resume by emailing Christy Edwards (Sr. e-Recruitment Consultant with
Personify) at ce-at-personifysearch.com. We wish everyone the best of luck.
Unfortunately only qualified candidates will be considered.

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10, 33 -- From: Christy Edwards {ce-at-personifysearch.com}
10, 33 -- MIME-Version: 1.0
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10, 33 -- Date: Mon, 26 Mar 2012 09:00:58 -0400
10, 33 -- Message-ID: {51a2676680ed269bbb98971f2b69f97b-at-mail.gmail.com}
10, 33 -- Subject: Fluorescence - Research Microscopy Specialist Opportunity
10, 33 -- To: Microscopy-at-microscopy.com
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==============================End of - Headers==============================

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

The Company:
Our client is a leading developer and producer of innovative high-tech
precision optics systems for the analysis of microstructures.� As one of
the market leaders in each of the fields of Microscopy, Confocal Laser
Scanning Microscopy, Imaging Systems, Specimen Preparation and Medical
Equipment.� Comprising nine manufacturing facilities in seven countries,
sales and service companies in 20 countries and an international network
of dealers, the company is represented in over 100 countries.

The Opportunity:
The company currently has an opening for a Microscopy Sales Representative
to be based in New York.� All applicants must not be adverse to travel, as
this is a position that may require you to travel when necessary.

Base: Commensurate with experience

Other: Full benefits - 401k program/matching - Car allowance - Company
cell phone - Laptop computer - Home office set-up - Expenses

Primary Responsibilities:
As a key member of the sales team, the purpose of this position is to
promote sales of the company's product line in the assigned region.� The
primary function is to execute domestic sales plans, recommend strategies
to improve the sale of products in the territory and aid in the
implementation of these strategies in line with Marketing, Corporate
growth, profitability and mission objectives.

Additional Responsibilities:
- Schedule product seminars or training sessions in concert with company
Product Management in the assigned region
- Participate in all activities that will enhance the awareness,
acceptance and sale of products
- Conduct analysis and performance evaluation of all sales by use of the
company's Customer Resource Management tool
- Work with Sales Manager to develop strategies designed to increase sales
of products; Specifically, the Microscopy Sales Representative will
recommend product, pricing, advertising and sales promotion strategies
- Coordinates the generation of new product ideas or modifications,
performs analysis and preliminary screening of these ideas and recommends
the appropriate ones to the Area Sales Manager for acceptance or rejection

Education and Experience Required:
BA/BS or MS degree in physics or chemistry or a related scientific field
is highly preferred.� Experience working with microscopy products is
required and a minimum of 2 years related microscopy experience in a
laboratory is essential.� In-depth knowledge of the microscopy market is
also highly desirable.� Sales experience in selling products of a
technical nature is desirable but not mandatory.� The individual must be
highly numerate as pricing, quoting and reviewing commercial terms will be
a daily routine. Strong computer skills and experience with CRM systems
(Maximizer, ACT, Goldmine, Siebel, etc).� Strong technical,
electrochemical, optical and mechanical aptitude.

If you meet and/or exceed the experience criteria, please submit your
resume by emailing Christy Edwards (Sr. e-Recruitment Consultant with
Personify) at ce-at-personifysearch.com. We wish everyone the best of luck.
Unfortunately only qualified candidates will be considered.

==============================Original Headers==============================
10, 33 -- From ce-at-personifysearch.com Mon Mar 26 08:58:02 2012
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10, 33 -- MIME-Version: 1.0
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10, 33 -- Message-ID: {2f782b6ec247d502cf2b12bc9b864048-at-mail.gmail.com}
10, 33 -- Subject: Microscopy Sales Representative - New York
10, 33 -- To: Microscopy-at-microscopy.com
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==============================End of - Headers==============================

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

The Company:
Our client is a leading developer and producer of innovative high-tech
precision optics systems for the analysis of microstructures.� As one of
the market leaders in each of the fields of Microscopy, Confocal Laser
Scanning Microscopy, Imaging Systems, Specimen Preparation and Medical
Equipment.� Comprising nine manufacturing facilities in seven countries,
sales and service companies in 20 countries and an international network
of dealers, the company is represented in over 100 countries.

The Opportunity:
The company currently has an opening for a Microscopy Sales Representative
to be based in Florida.� All applicants must not be adverse to travel, as
this is a position that may require you to travel when necessary.

Base: Commensurate with experience

Other: Full benefits - 401k program/matching - Car allowance - Company
cell phone - Laptop computer - Home office set-up - Expenses

Primary Responsibilities:
As a key member of the sales team, the purpose of this position is to
promote sales of the company's product line in the assigned region.� The
primary function is to execute domestic sales plans, recommend strategies
to improve the sale of products in the territory and aid in the
implementation of these strategies in line with Marketing, Corporate
growth, profitability and mission objectives.

Additional Responsibilities:
- Schedule product seminars or training sessions in concert with company
Product Management in the assigned region
- Participate in all activities that will enhance the awareness,
acceptance and sale of products
- Conduct analysis and performance evaluation of all sales by use of the
company's Customer Resource Management tool
- Work with Sales Manager to develop strategies designed to increase sales
of products; Specifically, the Microscopy Sales Representative will
recommend product, pricing, advertising and sales promotion strategies
- Coordinates the generation of new product ideas or modifications,
performs analysis and preliminary screening of these ideas and recommends
the appropriate ones to the Area Sales Manager for acceptance or rejection

Education and Experience Required:
BA/BS or MS degree in physics or chemistry or a related scientific field
is highly preferred.� Experience working with microscopy products is
required and a minimum of 2 years related microscopy experience in a
laboratory is essential.� In-depth knowledge of the microscopy market is
also highly desirable.� Sales experience in selling products of a
technical nature is desirable but not mandatory.� The individual must be
highly numerate as pricing, quoting and reviewing commercial terms will be
a daily routine. Strong computer skills and experience with CRM systems
(Maximizer, ACT, Goldmine, Siebel, etc).� Strong technical,
electrochemical, optical and mechanical aptitude.

If you meet and/or exceed the experience criteria, please submit your
resume by emailing Christy Edwards (Sr. e-Recruitment Consultant with
Personify) at ce-at-personifysearch.com. We wish everyone the best of luck.
Unfortunately only qualified candidates will be considered.

==============================Original Headers==============================
10, 33 -- From ce-at-personifysearch.com Mon Mar 26 08:58:29 2012
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10, 33 -- Subject: Microscopy Sales Representative - Florida
10, 33 -- To: Microscopy-at-microscopy.com
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==============================End of - Headers==============================

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

On Mar 26, 2012, at 8:27 AM, microscopylistserver-
noreply-at-microscopy.com wrote:

} Organization: Florida State University
}
} Title-Subject: [Filtered] TEM fee charge rate for internal and
} external
} academic users
}
} Message: I have a question concerning the TEM user fee charge rate for
} internal and external academic users. I noticed the common partice is
} that the internal academic users are charged at a lower rate than the
} external academic user. This sounds reasonable.
}
} However, I am told by my administrative people that we are not allowed
} to charge federally funded users rates that include unallowable costs.
} Whatever it means we are now charging the same rate, which does not
} sound right to me.
}
} I would appreciate if anyone can give me some information/rationale on
} the different academic rate.
}
} Thanks
} Yan Xin
} NHMFL/FSU

Dear Yan,
When I was working for New York State, we were by law not allowed
either to make a profit or to lose money from outside users, so
someone above my pay grade had to calculate the exact hourly cost of
running the facility. I don't know whether this is the case in
Florida. In any event, all outside users had to pay the same amount;
however, since we were an NIH Regional Resource, users with NIH grants
were not charged anything. Sometimes the charging structures can be
pretty complicated and not altogether rational.
Yours,
Bill

==============================Original Headers==============================
8, 38 -- From wtivol-at-sbcglobal.net Mon Mar 26 14:12:51 2012
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8, 38 -- To: microscopy-at-microscopy.com, xin-at-magnet.fsu.edu
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8, 38 -- Subject: Re: [Microscopy] viaWWW:TEM fee charge rate for internal and external academic users
8, 38 -- Date: Mon, 26 Mar 2012 12:12:48 -0700
8, 38 -- References: {201203261527.q2QFRPIo031725-at-ns.microscopy.com}
8, 38 -- X-Mailer: Apple Mail (2.936)
==============================End of - Headers==============================

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Yan,

Setting fair rates is indeed difficult for a university core facility.
Different institutions have slightly different policies but a few things
need to be considered:

1) The federal granting agencies do require that the rates are the same for
all those who have federal grants and that these are equal to those paid
by others at the institution. This means that you cannot allow some users
to pay less than others. Sometimes there is a desire to give new users a
break or arrange other incentives but this is not acceptable if it means
that others with NIH or other grants pay more.

2) External non-profit users are normally charged the same base rates as
internal users with the following exceptions:
They normally must pay the indirect charges that each institution also
deducts from grants received by internal researchers. These funds go to
maintain the institution facilities and pay for heat, electricity, etc
needed to run the facilities.

Often labor is subsidized for internal users in one way or another.
External users usually have to pay full salary + fringe benefit costs for
the labor associated with their using the institution facilities.

3) External for-profit users pay all these costs plus additional. Since the
university is not-for-profit, they must not charge less than commercial
sources for similar activities. Thus rates for external for-profit users
are normally significantly higher than those for internal or external
not-for-profit. You need to get an idea of external rates from other
institutions and commercial labs in your area so that you can justify your
rates.

Hope this helps,
Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu

Debra Sherman, Chief Scientific Officer
DS imaging, LLC
E-mail: debby.sherman-at-dsimagingllc.com
www.dsimagingllc.com

On 3/26/12 3:14 PM, "wtivol-at-sbcglobal.net" {wtivol-at-sbcglobal.net} wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
}
} On Mar 26, 2012, at 8:27 AM, microscopylistserver-
} noreply-at-microscopy.com wrote:
}
} } Organization: Florida State University
} }
} } Title-Subject: [Filtered] TEM fee charge rate for internal and
} } external
} } academic users
} }
} } Message: I have a question concerning the TEM user fee charge rate for
} } internal and external academic users. I noticed the common partice is
} } that the internal academic users are charged at a lower rate than the
} } external academic user. This sounds reasonable.
} }
} } However, I am told by my administrative people that we are not allowed
} } to charge federally funded users rates that include unallowable costs.
} } Whatever it means we are now charging the same rate, which does not
} } sound right to me.
} }
} } I would appreciate if anyone can give me some information/rationale on
} } the different academic rate.
} }
} } Thanks
} } Yan Xin
} } NHMFL/FSU
}
}
} Dear Yan,
} When I was working for New York State, we were by law not allowed
} either to make a profit or to lose money from outside users, so
} someone above my pay grade had to calculate the exact hourly cost of
} running the facility. I don't know whether this is the case in
} Florida. In any event, all outside users had to pay the same amount;
} however, since we were an NIH Regional Resource, users with NIH grants
} were not charged anything. Sometimes the charging structures can be
} pretty complicated and not altogether rational.
} Yours,
} Bill
}
}
}
}
} ==============================Original Headers==============================
} 8, 38 -- From wtivol-at-sbcglobal.net Mon Mar 26 14:12:51 2012
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18, 31 -- From dsherman-at-purdue.edu Mon Mar 26 21:13:18 2012
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Hi Everyone,

I have been asked to post this advert.

Imaging Facility Manager post

The Weatherall Institute of Molecular Medicine, University of Oxford is
seeking an Imaging Facility Manager to develop and manage the new state
of the art core facility. This new exciting role is funded for 5 years
in the first instance. To apply and for further details, including a job
description and selection criteria, please go to:

https://www.recruit.ox.ac.uk/pls/hrisliverecruit/erq_jobspec_version_4.jobspec?p_id=102426 .

Thanks,

Ian
--
Ian Dobbie
Micron Imaging Unit Manager,
Biochemistry,
University of Oxford,
South Parks Road,
Oxford
OX1 3QU
Tel: 01865 613323
Email: ian.dobbie-at-bioch.ox.ac.uk

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Dear Yan,

If I understand you correctly, your administrator has directed you to charge both external and internal users the same fees. Unless there has been a change in Federal policy, the only injunction applies to fees - for the use of instruments purchased/supported(?) with Federal funds - that unfairly complete with private businesses in the local region. Where there is no such competition, there can be no issue.

I have always counseled - to those who care - that there are some applications of microscopy in which I am not interested such as quality control. Thus, I have been willing to train operators from a private external corporation who can then pay an expanded hourly rate that does not unfairly compete with similar charges of private suppliers of microscopy services. If I managed a facility at the University of Chicago and wanted to open the facility to 'work' sent from private corporations for microscopy services I would have the following considerations (admittedly somewhat complicated) to prevent me from unfairly competing with McCrone Associates capabilities (http://www.mccroneassociates.com/techniques ).
1. I could charge fees exactly like McCrone's for similar services.
2. I could offer to McCrone as to any other organization access to those instrumentalities that are not included in the McCrone possessions that I happen to have in my facility. I could refer to McCrone as McCrone could refer to me - very collegial!
3. I can charge internal users whatever the financial administrators of the University of Chicago felt would be accepted by the next funding agency reviewers who will fund charges for use by 'fundees', but will generally NOT fund requests for lump sum service contracts. This work is for institutional bean counters and the cadre of users who work in the institutions and win the grant money administered by them.

My point is that Florida State, NOT Yan Xin, can adequately determine its potential for breaching the 'intentions' of the Federal regulations. Policies that I like to apply, must be approved by superiors. Indeed, when I speak to superiors about these issues, I always try to bring a small stack of papers that correctly illuminate the Federal Regulation and what I consider to be viable options for the facility I manage - which, by the way, has never been at an institution with large amounts of Federal funding.

If you wish, you may contact me privately for copies of those documents that I carry with me to meetings with admins who 'run the joint'. If you are going to manage, my policy is to go informed.

My first fee schedule was derived from those I could take from facilities at institutions that had large amounts of Federal funding. My rationale was that such fees would generally reflect fees that would be acceptable to the Federal funding agencies like NIH and NSF.

Hope this helps

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

If one looks long enough, one may actually see something frozen in time that flashes one's mind to a new state.
1. The day that I asked myself how the hydrogen atom's mass could actually be: 1.0008.
2. The day, while looking at a slide of some tissue, I saw a leukocyte that had been caught sneaking in or out of a capillary: http://www.annualreviews.org/doi/abs/10.1146/annurev-pathol-011110-130224

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Email: xin-at-magnet.fsu.edu Name: Yan Xin

Organization: Florida State University

Title-Subject: [Filtered] TEM fee charge rate for internal and external academic users

Message: I have a question concerning the TEM user fee charge rate for internal and external academic users. I noticed the common partice is that the internal academic users are charged at a lower rate than the external academic user. This sounds reasonable.

However, I am told by my administrative people that we are not allowed to charge federally funded users rates that include unallowable costs.
Whatever it means we are now charging the same rate, which does not sound right to me.

I would appreciate if anyone can give me some information/rationale on the different academic rate.

Thanks
Yan Xin
NHMFL/FSU

Login Host: 146.201.233.77
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Fellow Microscopists,

I am finding the need for a new lab camera to replace one I gave away and I was hoping for some suggestions from the group as to what I should purchase.

To start with, I need to be able to take images of mounts, like SEM stubs and carbon planchets, so it needs the capacity to take clear macro images. There are some times when attaching the camera to a stand just won't work or is awkward, so good image stabilization is a plus, so I can hold it and still get good images. The lab also has a microscope, a Leica MZ-6, that has no camera or camera adapter, so I would like to use this camera for the scope if possible. There is no capacity to keep and maintain a computer in the lab, which is a cleanroom, so it has to be able to run on its own without a computer. I am hoping to remove the memory card and recover images in a different building.

Thanks in advance for any advice.

Abby

Abigail P. Lindstrom

NIST-MML-637.05
100 Bureau Drive
MS 8371
Gaithersburg, MD 20899-8371

301 975-5172
fax 301 417-1321

==============================Original Headers==============================
9, 24 -- From abigail.lindstrom-at-nist.gov Tue Mar 27 10:04:37 2012
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9, 24 -- From: "Lindstrom, Abigail P." {abigail.lindstrom-at-nist.gov}
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9, 24 -- Date: Tue, 27 Mar 2012 11:04:35 -0400
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Try holding an iPhone camera about an inch above the eyepiece of your microscope. With a little practice, the results will surprise you.

John Mardinly
ASU

On Mar 27, 2012, at 8:12 AM, "abigail.lindstrom-at-nist.gov" {abigail.lindstrom-at-nist.gov} wrote:

}
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} Fellow Microscopists,
}
} I am finding the need for a new lab camera to replace one I gave away and I was hoping for some suggestions from the group as to what I should purchase.
}
} To start with, I need to be able to take images of mounts, like SEM stubs and carbon planchets, so it needs the capacity to take clear macro images. There are some times when attaching the camera to a stand just won't work or is awkward, so good image stabilization is a plus, so I can hold it and still get good images. The lab also has a microscope, a Leica MZ-6, that has no camera or camera adapter, so I would like to use this camera for the scope if possible. There is no capacity to keep and maintain a computer in the lab, which is a cleanroom, so it has to be able to run on its own without a computer. I am hoping to remove the memory card and recover images in a different building.
}
} Thanks in advance for any advice.
}
} Abby
}
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HI ABIGALE..

YOU HAVE AN MZ6, NOT REALY SUITABLE FOR A TRINOC FITTING, WHY NOT ADD AN OCCULAR
CAMERA (DIGITAL) AND RUN A USB OUT OF YOUR CLEAN ROOM INTO YOUR PC, FAILING THAT
IT IS POSS TO BUY AN OCULAR CAMERA WITH MEMORY CARD INSIDE THE CAMERA.

I SUGGEST BRUNEL MICROSCOPES IN UK, I HAVE NEVER FOUND ANYONE BETTER.

HOPE YOU FIND WHAT SUITS YOU.

DAVID BRADBURY..UK

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Yan's original question was about fees for internal vs external federal
funded academic users, It had nothing to do with fees for federal funded
vs private users. Our fees for external academic users and internal users
are same. One intention (among others) for such a practice is to
encourage research collaborations. As far as I know most university EM
cores do charge private industry users at a higher rate.

Hong

On 3/27/12 10:57 AM, "FMonson-at-wcupa.edu" {FMonson-at-wcupa.edu} wrote:

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9, 41 -- From hyi-at-emory.edu Tue Mar 27 11:12:40 2012
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Hi Ravi,

Small image shifts may occur due to changes in the intermediate and projector lenses. Any TEM built after about 1990 should be able to be aligned to have minimal image shift with magnification changes. Call your local service provider and ask for a full column alignment. Once done, it is good for years--assuming your batteries are working that back up your alignment data in the TEM.

Best regards,
Roseann

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

On Mar 27, 2012, at 6:15 AM, microscopylistserver-noreply-at-microscopy.com wrote:

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} Email: ravi.thakkar369-at-gmail.com Name: Ravi Thakkar
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} Organization: National Institute of Mental Health And Neuro Science
}
} Title-Subject: [Filtered] TEM : Image shift during changing the perticular magnification.
}
} Message: Dear Listeners, The Image is shifting during changing the particular magnification, not for
} the all of the magnification.
}
} Is this related to changing the current in lens objective or projector lens system..??
}
} Kindly explain me in brief.
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} Login Host: 14.139.159.99

==============================Original Headers==============================
9, 44 -- From rcsencsits-at-lbl.gov Tue Mar 27 11:20:47 2012
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9, 44 -- From: Roseann Csencsits {rcsencsits-at-lbl.gov}
9, 44 -- Subject: Re:Image shift during changing the particular magnification
9, 44 -- Date: Tue, 27 Mar 2012 09:20:42 -0700
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Dear Abby,

There is a range of adapters you can get to attach your regular digital
camera to a microscope via one of the eyepiece slots - usually you need to
take the eyepiece out and put the adapter in. That's definitely the best
way in my opinion.
Re. image stabilisation - nothing like going and testing a few cameras
yourself in the kind of lighting you expect to be using it in.
cheers,
Rosemary

Dr Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
E rosemary.white-at-csiro.au

On 28/03/12 2:08 AM, "abigail.lindstrom-at-nist.gov"
{abigail.lindstrom-at-nist.gov} wrote:

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Colleagues....

Just testing, I have a small problem on the Listserver that
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Your question is rather timely. I had someone asking to look at some samples of concrete yesterday. Like them, I wonder if your researcher has thought things through very well.

What do they want to see? How do they expect to do so? What kind of preparation and magnification will be required? Those issues may pose bigger issues.

Our first attempt to look at concrete in an SEM goes back almost 20 years. We prepared a piece between 1 and 2 inches square and about 1/4 inch thick, coated it with gold, and placed it into the SEM and started pumping. Our JEOL 840A gave up after some time (30 minutes?) as it was not able to move beyond the roughing cycle. We were able to bypass the timer and left the SEM pumping over night. It had reached working vacuum by the next morning. However, examination showed extensive cracking where the vacuum had pulled water out of the structure. (I don't recall how completely cured it was.)

Therefore, high-vacuum characterization was not an option. We purchased an SEM with variable pressure capabilities and have used it for over 15 years. Most of our work is done at 40 Pa (~0.3 Torr). That means we achieve operating vacuum quickly, the residual atmosphere alleviates charging, and samples are not grossly changed by exposure to high vacuum.

You definitely have a non-conductive sample that either needs coating or needs to be examined in V-P mode. I don't know that asphalt samples would have such troubles with out-gassing, but I expect they would. A sample might seem stable at room temperature and pressure, but the lower pressure of an SEM would extract additional volatile components. That would change the nature of your sample and present pumping and possible contamination issues. The binder would heat some under the beam and that would release more components. As the binder loses components, I would expect any conductive metal layer to be disrupted and the sample would start charging. A cold finger might help capture those components so they don't condense somewhere bad. Your cold finger would then need cleaning.

One significant benefit of V-P mode is that the residual gas changes the pumping regime. It helps to sweep contaminants out of the chamber rather than letting them diffuse throughout and condense on surfaces.

For those reasons, I would not try to examine the sample in high-vacuum. I might let them try to examine a small piece, but I expect the examination would be a failure because of changes to the sample. I would try V-P mode, but I would watch for changes in the sample to see if you are truly seeing the sample as it is.

That brings me back to the questions of what they want to see and how they will prepare the sample.

If they simply grab a blob of asphalt concrete, they will see the aggregate shapes and should be able to see the gross structure, including large-scale porosity. However, if they zoom in on the aggregate, they will see most, if not all, of the aggregate coated with binder. The beam will have difficulty penetrating through the binder to the aggregate. A stereo microscope could do about as well.

If they try to prepare a section through the sample, they will be able to discern things at low magnification. I would be concerned about the binder migrating over the surface and causing difficulties at high magnification. Again, a stereo microscope or reflected light microscope might do as well.

So I would ask again, why are they considering SEM, what do they hope to learn or document, and how? I encounter many students who really do not consider those questions before bringing me their samples. Help them to bring those issues into focus.

Warren Straszheim

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Title-Subject: [Filtered] Asphalt in high vac SEM

Message: Hello everybody,

A researcher wants to examine solid asphalt specimens under out high-vacuum SEM. Is it safe to the
SEM. Would that contaminate the column or the TMP?

Thank you very much..

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Ahmad,
Unless you have a cold stage (and use a very small sample on that stage),
you are probably going to contaminate your system.

Is that a problem? It depends on how the system has been used in the past
and how you intend to use it in the future. If you are running LaB6 or it
has a field emission gun I would not put the asphalt in the system without a
cold stage. On the other hand, if it is an old tungsten gun and the TMP is
brought to atmosphere with every specimen change, you may not notice any
additional contamination.

You can also lessen the amount of contamination by operating at a low kV and
small spot size (high condenser lens current) to minimize heating. If you
need to do x-ray analysis, you are going to heat the sample. Period.

If the system has been kept very clean and quantitative analysis is a major
part of its job, I would not put asphalt in. On the other hand, if it is an
old system and no one has been particularly careful about what goes in, and
it's mostly used for imaging and maybe some qualitative analysis, then I
might consider it.

To actually answer your question, yes, it is going to contaminate your
vacuum system unless you have a very small sample on a cold stage. If the
system is currently very clean, then yes, it would be considered damaging to
the system.

Good luck!

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Title-Subject: [Filtered] Asphalt in high vac SEM

Message: Hello everybody,

A researcher wants to examine solid asphalt specimens under out high-vacuum
SEM. Is it safe to the
SEM. Would that contaminate the column or the TMP?

Thank you very much..

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Detector is a PGT Prism from circa 1997. Condensation / ice forms on probe when specimen chamber is opened; re-running auto cal works but only for a while, clearly something (vacuum?) is wrong with the detector.

SEM is ISI DS-130 from 1986, runs perfectly. Pulse processor / software is IXRF from 2007 or so. Most work is metallurgical failure analysis, some geological samples, some well debris (anything you might find downhole in an oil or gas well could show up in my lab).

I recall reading in the Microscopy Listserv about someone named Jim Nicolino (sp? - maybe do not recall his name correctly, am bad with names but this one stuck in my mind long ago since I thought I might need the detector rebuilt at some point) who was reputed to do excellent work.

At this point I'm pondering rebuilding versus replacing. Personally, I dislike "buying a new on" (my pickup has over 488k miles on it) but the SEM/EDS is not "mine", it belongs to the company, and I owe it to them to choose wisely. I can and will educate myself on available "new" detectors if y'all tell me that is the way to go - this is the first step in that process.

So - respectfully requesting opinions and guidance, rebuild or replace (and with what)? Thanks in advance.

Regards,
Andrew T. Werner
Chief Metallurgist, Perforating
Schlumberger Reservoir Completions
14910 Airline Road, Rosharon, TX 77583-1590
(281) 285-5272

"I Shoot the hippopotamus with bullets made of platinum,
because if I use leaden ones his hide is sure to flatten 'em"
The Bad Child's Book of Beasts - Hilaire Belloc

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Warren,

I agree with all your statements but one:
"SEM with variable pressure capabilities ... at 40 Pa (~0.3 Torr). That means ... samples are not grossly changed by exposure to high vacuum."

Pressure 0.3 Torr is just 0.4% of atmospheric pressure. It means that compared to pressure at high vac mode 99.6% of pressure difference is still applied to the specimens. For my porous specimens (bone, teeth) I do not see any difference in crack development at high vac mode or at variable pressure. The same goes for water: only 0.4% of partial water pressure could be preserved at 0.3 Torr, so water would evaporate rather quickly, at about the same rate as in high vac mode.

I believe specimen protection from difference in pressures and/or any water preservation in specimen are not among benefits of variable pressure mode.

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: wesaia-at-iastate.edu [mailto:wesaia-at-iastate.edu]
} Sent: Wednesday, March 28, 2012 9:56 AM
} To: Dusevich, Vladimir
} Subject: [Microscopy] RE: viaWWW:Asphalt in high vac SEM
}
}
}
}
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}
} Your question is rather timely. I had someone asking to look at some samples
} of concrete yesterday. Like them, I wonder if your researcher has thought
} things through very well.
}
} What do they want to see? How do they expect to do so? What kind of
} preparation and magnification will be required? Those issues may pose
} bigger issues.
}
} Our first attempt to look at concrete in an SEM goes back almost 20 years. We
} prepared a piece between 1 and 2 inches square and about 1/4 inch thick,
} coated it with gold, and placed it into the SEM and started pumping. Our JEOL
} 840A gave up after some time (30 minutes?) as it was not able to move
} beyond the roughing cycle. We were able to bypass the timer and left the
} SEM pumping over night. It had reached working vacuum by the next
} morning. However, examination showed extensive cracking where the
} vacuum had pulled water out of the structure. (I don't recall how completely
} cured it was.)
}
} Therefore, high-vacuum characterization was not an option. We purchased
} an SEM with variable pressure capabilities and have used it for over 15 years.
} Most of our work is done at 40 Pa (~0.3 Torr). That means we achieve
} operating vacuum quickly, the residual atmosphere alleviates charging, and
} samples are not grossly changed by exposure to high vacuum.
}
} You definitely have a non-conductive sample that either needs coating or
} needs to be examined in V-P mode. I don't know that asphalt samples would
} have such troubles with out-gassing, but I expect they would. A sample
} might seem stable at room temperature and pressure, but the lower
} pressure of an SEM would extract additional volatile components. That would
} change the nature of your sample and present pumping and possible
} contamination issues. The binder would heat some under the beam and that
} would release more components. As the binder loses components, I would
} expect any conductive metal layer to be disrupted and the sample would
} start charging. A cold finger might help capture those components so they
} don't condense somewhere bad. Your cold finger would then need cleaning.
}
} One significant benefit of V-P mode is that the residual gas changes the
} pumping regime. It helps to sweep contaminants out of the chamber rather
} than letting them diffuse throughout and condense on surfaces.
}
} For those reasons, I would not try to examine the sample in high-vacuum. I
} might let them try to examine a small piece, but I expect the examination
} would be a failure because of changes to the sample. I would try V-P mode,
} but I would watch for changes in the sample to see if you are truly seeing the
} sample as it is.
}
} That brings me back to the questions of what they want to see and how they
} will prepare the sample.
}
} If they simply grab a blob of asphalt concrete, they will see the aggregate
} shapes and should be able to see the gross structure, including large-scale
} porosity. However, if they zoom in on the aggregate, they will see most, if
} not all, of the aggregate coated with binder. The beam will have difficulty
} penetrating through the binder to the aggregate. A stereo microscope could
} do about as well.
}
} If they try to prepare a section through the sample, they will be able to
} discern things at low magnification. I would be concerned about the binder
} migrating over the surface and causing difficulties at high magnification.
} Again, a stereo microscope or reflected light microscope might do as well.
}
} So I would ask again, why are they considering SEM, what do they hope to
} learn or document, and how? I encounter many students who really do not
} consider those questions before bringing me their samples. Help them to
} bring those issues into focus.
}
} Warren Straszheim
}
} -----Original Message-----
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} Subject: [Microscopy] viaWWW:Asphalt in high vac SEM
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} Email: ahmad_ashkaibi-at-hotmail.com Name: Ahmad Ashkaibi
}
} Organization: Al-Blaqa University
}
} Title-Subject: [Filtered] Asphalt in high vac SEM
}
} Message: Hello everybody,
}
} A researcher wants to examine solid asphalt specimens under out high-
} vacuum SEM. Is it safe to the SEM. Would that contaminate the column or
} the TMP?
}
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Hello everyone,
I wonder if anyone has or know someone or a group that has a SEM with
a heating stage. I would like to heat a sample up to 900 degree C and
image it before and after heating. The reason I can not do the heating
in a tube furnace under an inert gas, for example, is the the sample
is very sensitive to oxygen a high temperature and the heat treatment
will have to be done under vacuum,which is something we do not have in
our group.

If there is a high vacuum chamber with a heating stage that can go up
to 900 degree C, this will be great.

Any help is very appreciated

Thank you

--
**********************************************************
Hany El-Sayed, Ph.D.
Post-doctoral�Fellow
Department of Chemistry
University of Calgary, Alberta
Canada
403-923-7802
helsayed-at-ucalgary.ca
**********************************************************

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Good morning,
Dear Andrew,

you perhaps ment this one :
(personal note: the website given below still is active/accessible )

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e-mail by NWWhite-at-bwxt.com Thursday Do 18.10.2007 19:54

[Microscopy] RE: Looking for help repairing an EDS detector

Hello Chris,

It was several years ago, but (I) Jim Nicolino satisfactorily rebuilt one
of my LN2 EDS detectors. No vested interest.
Latest web site I have:

http://advancedanalysistech.com/index2.html

Woody White
BWXT Services

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=====================================================================================================
Best wishes, HAPPY EASTER, good luck and
warm regards

Wolfgang MUSS
Salzburg, Austria

=================================
+++ 39th Ann. SCUR (Society for Cutaneous Ultrastructure Research) Meeting LYON, France +++
23-25 May, 2012, informations see:
http://orgs.dermis.net/content/e04scur/e03meetings/e770/e771/index_ger.html

Forthcoming in 2013:
+++ 40th Ann. SCUR Meeting SALZBURG, Austria/Autriche +++
Joint Meeting with the SSSR (Society for Skin Structure Research, Japan)
12-14 May 2013, further informations following ASAP
see also
http://orgs.dermis.net/content/e04scur/e03meetings/e770/e771/index_ger.html

=====================================================================================================

} -----Urspr�ngliche Nachricht-----
} Von: werner1-at-slb.com [mailto:werner1-at-slb.com]
} Gesendet: Mittwoch, 28. M�rz 2012 18:15
} An: Mu� Wolfgang
} Betreff: [Microscopy] EDS detector - rebuild or replace?
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} Detector is a PGT Prism from circa 1997.
} Condensation / ice forms on probe when specimen chamber is opened; re-
} running auto cal works but only for a while, clearly something
} (vacuum?) is wrong with the detector.
}
} SEM is ISI DS-130 from 1986, runs perfectly.
} Pulse processor / software is IXRF from 2007 or so.
} Most work is metallurgical failure analysis, some geological samples,
} some well debris (anything you might find downhole in an oil or gas
} well could show up in my lab).
}
} I recall reading in the Microscopy Listserv about someone named Jim
} Nicolino (sp? - maybe do not recall his name correctly, am bad with
} names but this one stuck in my mind long ago since I thought I might
} need the detector rebuilt at some point) who was reputed to do
} excellent work.
}
} At this point I'm pondering rebuilding versus replacing. Personally, I
} dislike "buying a new on" (my pickup has over 488k miles on it) but the
} SEM/EDS is not "mine", it belongs to the company, and I owe it to them
} to choose wisely. I can and will educate myself on available "new"
} detectors if y'all tell me that is the way to go - this is the first
} step in that process.
}
} So - respectfully requesting opinions and guidance, rebuild or replace
} (and with what)? Thanks in advance.
}
} Regards,
} Andrew T. Werner
} Chief Metallurgist, Perforating
} Schlumberger Reservoir Completions
} 14910 Airline Road, Rosharon, TX 77583-1590
} (281) 285-5272
}
} "I Shoot the hippopotamus with bullets made of platinum,
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Thank you all for your very helpful comments!

I will reply individually, but right now it looks like Doug Connors, to whom Jim Nicolino sold his business when he retired, is going to bake out and re-evacuate the detector - for a relatively small fraction of the cost of a rebuild or replacement. His diagnosis, based on the fact that the detector works fine except for forming ice and needing frequent recalibration, is that the vacuum has simply degraded over 15 years of use.

Thank you again! This List is a marvelous resource.

Regards,
Andrew

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South Central Texas, Austin area

Brief Description & Qualifications:
Newly created role to support increased demand in the U.S.A for the
microscopy division of global analytical analysis.
Seeking a talented, practical technologist/engineer possessing 3
years+ "best practice" industry exposure in technical support within
advanced microscopy (TEM/SEM/EDS/EBSD/WDS/XRF)

This professional is going to work alongside other team members in
applications, installation, training, and technical service/support
environments.
BS Engineering/Physics and Technical Masters desirable (industry
experience may offset).
Ability and desire to travel 25%-40%. �Must be comfortable working
independently at
customer sites.

Please send cover letter and resume to:� microscopyhire-at-gmail.com

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All,

A scientist has asked to image porcine virus in our FESEM. She says that it is listed as a biosafety level 2, but has been disinfected to } 99% level with 10% glutaraldehyde. Are any of you working with this virus? I'm not familiar with the safety protocols. Is this a safe project for a core facility?

Thanks in advance.

Owen

==============================Original Headers==============================
7, 20 -- From prvs=4299dd50d=opmills-at-mtu.edu Fri Mar 30 09:45:05 2012
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It isn't really up to you or the PI to decide if this is safe to do. Your university undoubtedly has a biosafety committee. Only they can approve the use of a BSL-2 pathogen in your core facility. The PI needs to have a protocol in place that has been approved by the institutional committee.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: opmills-at-mtu.edu [mailto:opmills-at-mtu.edu]
Sent: Friday, March 30, 2012 9:46 AM
To: Phillips, Thomas E.

All,

A scientist has asked to image porcine virus in our FESEM. She says that it is listed as a biosafety level 2, but has been disinfected to } 99% level with 10% glutaraldehyde. Are any of you working with this virus? I'm not familiar with the safety protocols. Is this a safe project for a core facility?

Thanks in advance.

Owen

==============================Original Headers==============================
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Greetings Fellow Microscopists,

The Midwest Microscopy and Microanalysis Society will hold its next meeting on Thursday, April 26th, at the University of Illinois at Chicago. Robert Klie and Ke-Bin Low have lined up an impressive group of speakers for this program, "Seeing is Believing: Aberration Corrected STEM". Attendees will also have the opportunity to view the recently installed JEOL JEM-ARM200CF in the Research Resources Center at UIC. Details can be found on our website under Meetings:

http://www.midwestmicroscopy.org/

Please join us to hear a great lineup of talks, visit with exhibitors and mingle with your local microscopy colleagues. We look forward to seeing you there!

Elaine Schumacher
M3S Program Coordinator

*********************************************************************
Elaine F.Schumacher
Senior Research Scientist
McCrone Associates, Inc.
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7417 (fax)
E-mail: eschumacher-at-mccrone.com
Web Site: www.mccrone.com

*********************************************************************
This message and any attachments are solely for the
intended recipient. If you are not the intended recipient,
disclosure, copying, use or distribution of the information
included in this message is prohibited.
*********************************************************************

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Dear Owen,

unless it is true and {verified} (whatever one means when saying "fixed") that the virus (suspension, particles, precipitate or whatsoever) previously has been } classically { fixed (conformal change of protein structures e.g. by suited aldehydes) [= deactivated by '10% glutaraldehyde-solution' I don't think the virus particles to be infectious any more so IMHO these haven't to be included under BSL-2 pathogen strategy (if such deactivated particles aren't included too in the agenda of a biosafety committee).
There are some papers concerning deactivations for personal as well as working safety by e.g. 1-2% FA and / or 1-2% GA, so the question rather is whether you'll see or can image something of "fine" structure of virus particles after 10% GA (only diluted?, buffered?, incubation time for deactivation?).
The ultrastructural aspect of virus particles deactivated that way has been reported to be of poor(er) quality than images taken after classical negative staining.

For example:
- (2007)considerations of RKI (Robert Koch Institute Berlin)-Regulations....
"Formaldehyde alone not necessarily inactivates virus particles, whereas addition of at least 0.5% Glutaraldehyde seems to inactivate properly"

I also would like to refer you to recent Listserver threads (see Archives):

{ {BSL-2 question (Nov. 3, 2009) by Paula Sicurello (vapatpxs-at-yahoo.com)
or

{ {TEM Negative staining HIV particles, a safety question (Nov 25,2011) by 2011 (hchen3-at-unl.edu)
} } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } }
in this thread, ==} [Microscopy] Re: TEM Negative staining HIV particles, a safety question.
sent by: paul_hazelton-at-umanitoba.ca Friday / Fr 25.11.2011 15:18 he says:
{Han
To my knowledge all studies ever conducted has shown that HIV is inactivated by standard fixation methods.
In fact, fixation as low as 0.2% Glutaraldehyde is considered to be effective.

paul hazelton
--
Paul R. Hazelton, PhD
Department of Medical Microbiology
Head of Electron Microscopy
Lab Director, Viral Gastroenteritis Study Group
University of Manitoba
511 Basic Medical Sciences Building
745 Bannatyne Avenue
Winnipeg, Manitoba, Canada, R3E 3J7
e-mail: paul_hazelton-at-umanitoba.ca
paulhazelton-at-mts.net
Phone: 204-789-3313 (w);
204-489-6924 (h)
Cell: 204-781-6982
Fax: 204-789-3926
} } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } } }

BUT, YES: Nevertheless, in the same thread, not to "hide" that:
} } [Microscopy] Re: TEM Negative staining HIV particles, a safety question. == General Aspects Biosafety==
Sent by: PWebster-at-hei.org Friday / Fr 25.11.2011 20:02Paul Webster said:

(Note added: "live/vivid HIV particles")
Dear Han (and everyone else),
Biosafety issues are not a subject for casual discussion on a listserver, they are to be addressed by the relevant institutional biosafety committee.
For Han, I suggest he stop the researcher using the HIV until an approved IBC protocol is in place. This protocol should address how the virus will be handled before it gets to the EM lab, how it will be handled in the EM lab and how it will be safely disposed of once it has been imaged.
The protocol should address suitable labeling of all materials to be used in the lab, warning signs outside and inside the lab, and relevant personal protective equipment.
The biosafety committee is a group of experts who will understand the safety issues of handling biological hazards and will offer the best advice for handling the virus.
The other users have every right to be concerned by potential exposure to virus until there is an approved IBC protocol in place.
The protocol should also address issues such as how the TEM specimen holder will be sterilized after use so that it can be safely handled by the other users. Again, the safety issue itself may not be important (if the virus is chemically fixed and has been irradiated), but the safety of all users has to be addressed first.
An approved IBC protocol should be in place for any lab that is handing viral, bacterial and fungal pathogens: for human cell lines and human bodily fluids. If the lab is routinely handling unfixed human materials it is also advisable for the staff to be vaccinated against the hepatitis b virus.
Failure to adhere to approved protocols (or failure to even apply for them) can result in loss of NIH funding and law suits.

Regards,

Paul Webster.

House Research Institute
2100 W. 3rd St.
Los Angeles,
CA 90057, USA.

I thought this was so important an issue that I am willing to bear the Thanksgiving "out of office" junk messages that await - does anyone unsubscribe when the go away from the office? { {
===================

Hope this to be of help,

best wishes and regards,
HAPPY EASTER-Time to you all,

Wolfgang MUSS PhD
Member of MSA
SALZBURG, AUSTRIA

} -----Urspr�ngliche Nachricht-----
} Von: opmills-at-mtu.edu [mailto:opmills-at-mtu.edu]
} Gesendet: Freitag, 30. M�rz 2012 16:49
} An: Mu� Wolfgang
} Betreff: [Microscopy] Porcine virus
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All,
A scientist has asked to image porcine virus in our FESEM.

She says that it is listed as a biosafety level 2, but has been disinfected to } 99% level with 10% glutaraldehyde. Are any of you working with this virus?

I'm not familiar with the safety protocols. Is this a safe project for a core facility?

Thanks in advance.

Owen
}
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In order to properly serve the needs of the growing materials science and engineering research community at Mississippi State University (MSU), the Institute for Imaging & Analytical Technologies (I2AT) has an opening for a Postdoctoral Research Associate with expertise in Transmission Electron Microscopy techniques as applied to the area of physical metallurgy. This Postdoctoral Research Associate will report to the Director of the I2AT, a university-wide core facility and research institute meeting all three missions of the university, research, teaching, and service.

The applicant will be responsible for operation, training, consultation, scheduling, and routine maintenance of new state-of-the-art multi-user analytical transmission electron microscope within MSU's I2AT. The applicant will primarily support TEM applications, but will also have the opportunity to learn/operate other imaging and characterization technologies.

The successful applicant must have a Ph.D. in any engineering or physical science field and a strong background in materials characterization, imaging and diffraction techniques, EM theory and operation. Knowledge of other general imaging/analysis approaches is desired.

For more information or to apply, see the link below:
https://www.jobs.msstate.edu/applicants/jsp/shared/frameset/Frameset.jsp?time=1333134701254

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A postdoctoral position is available at the University of Alberta to
study the structural biology of the infectious prion protein with a
particular emphasis on electron microscopy.

The laboratory is situated within the Centre for Prions and Protein
Folding Diseases, a multi-disciplinary and cross-faculty facility
comprised of a transgenic rodent facility and large communal
containment labs for level 2+ prion work. The Centre houses seven
intramural researchers with collaborative interests in prion and
protein folding diseases and whose expertise crosses a wide range.
Trainees at the Centre are exposed to weekly seminars, data
discussions, and have many opportunities for collaborations with other
labs nationally and internationally.

Applicants must have a Ph.D. and proven experience in the field of
structural biology, preferentially including electron microscopy
and/or cryo-electron microscopy. Excellent oral and written skills and
the ability to collaborate with others are required.

For more information please visit the following websites:
http://www.prioncentre.ca/
http://biochem.med.ualberta.ca/Pages/default.aspx
http://biochem.med.ualberta.ca/Research/faculty/AssociateProfessors/Pages/Holger-Wille.aspx

Interested candidates should submit a CV, the names and addresses of
three references, and a letter stating research interests and
experience.

Holger Wille
Associate Professor
University of Alberta
Centre for Prions and Protein Folding Diseases
204 Environmental Engineering Building
Edmonton, AB T6G 2M8
Canada

e-mail: wille-at-ualberta.ca

Phone: (780) 248-1712
Fax: (780) 492-9352

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On behalf of Kirk Czymmek:

Dear Microscopists,

As part of a series of advanced microscopy Workshops at the University of
Delaware Bio-Imaging Center located in the Delaware Biotechnology Institute
(http://www.dbi.udel.edu/directions.html), we have two upcoming multi-day
workshops within the next month and sponsored by leading microscopy
manufacturers that may be of interest to you.

The Bio-AFM Workshop (Bruker) is in a few weeks (April 11 and 12) and will
include the opportunity for hands-on AFM imaging of your difficult samples
with the experts. In addition, we will have a high-speed AFM available. For
more details and instructions to register, please see the link below:

http://bioimaging.dbi.udel.edu/Events/Bruker_April_2012

Space is limited and the deadline is approaching so please be sure to
sign-up soon.

Our Cryo-SEM Workshop (with Gatan) will be held May 1-3, includes (2)
one-day informative workshops that will address numerous applications using
cryo-techniques in SEM. Cryo applications include biological samples such
as tissue, cells, hair and more; other applications include botany,
polymers and any other samples that aresensitive to the electron beam.
Besides presentations on the use andapplication of cryo-techniques, there
will be a live demonstration of the instrumentation with samples on a
Hitachi FE-SEM S4700 with a Gatan Alto 2500 cryo system. Please see link
below for registration details:

http://bioimaging.dbi.udel.edu/gatan-cryo-sem-workshop

We look forward to seeing you there. If you have any questions, please feel
free to email me. (kirk-at-udel.edu).

Best Regards, Kirk

Kirk J. Czymmek, Ph.D.

Associate Professor

Department of Biological Sciences

Director UD Bio-Imaging Center

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SENIOR TECHNICAL OFFICER � ATOM PROBE SPECIALIST
SYDNEY MICROSCOPY & MICROANALYSIS
THE UNIVERSITY OF SYDNEY
REFERENCE NO. 374/0312

� Provide specialist technical support of the atom probe facilities and various ancillary techniques
� Full-time continuing, remuneration package: $84K p.a. which includes leave loading and up to 17% super

Sydney Microscopy & Microanalysis (SMM) aims to be a world-leading facility for modern microscopy and microanalysis, offering premier instruments, services and training to researchers from across Sydney and around Australia, and providing strong leadership in the national and international characterisation communities.

Reporting to the Laboratory Manager, you will provide specialist technical support of the centre�s atom probe facilities, focussing on the needs of users in atom probe microscopy and atom probe tomography so that the facilities provide effective high-quality outcomes for users.

In this role you will:

� instruct, train and assist users in their research work using the centre�s instrumentation, specimen preparation and/or operation of the instrument and/or analysis of atom probe data
� work cooperatively and constructively with colleagues to facilitate microscopy experiments for users and provide laboratory and microscopy services to commercial clients of the centre
� use the centre�s IT systems in order to ensure the smooth running and storage of data
� recommend process improvements.

To qualify for this role, you will demonstrate:

� advanced problem-solving skills and demonstrated ability to run and maintain all operational aspects of atom probe microscopy instrumentation (viz. electrical, electronic, mechanical, pneumatic, and ultra-high vacuum systems and the computer systems)
� a capacity to master Cameca visualisation and simulation software
� demonstrated skills in the application and/or development of advanced microscopy and microanalysis techniques.

The successful candidate must possess a Bachelor�s degree in electronic engineering, materials engineering, mechanical engineering, nanotechnology or physics.

All applications must be submitted via the University of Sydney careers website.� Visit sydney.edu.au/positions and search by the reference number for more information and to apply.

CLOSING DATE: 10 April 2012, 1pm

The University is an Equal Opportunity employer committed to equity, diversity and social inclusion. Applications from equity target groups and women are encouraged.

� The University of Sydney

==============================Original Headers==============================
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PROFESSIONAL OFFICER - X-RAY MICROSCOPY
SYDNEY MICROSCOPY & MICROANALYSIS
THE UNIVERSITY OF SYDNEY
REFERENCE NO. 373/0312

. Coordinate X-ray microscopy and image analysis facilities and the various ancillary techniques to deliver high quality user support
. Full-time continuing, remuneration package: $103K p.a. which includes leave loading and up to 17% super

Sydney Microscopy & Microanalysis (SMM) aims to be a world-leading facility for modern microscopy and microanalysis, offering premier instruments, services and training to researchers from across Sydney and around Australia, and providing strong leadership in the national and international characterisation communities.

Reporting to the Laboratory Manager, you will coordinate the centre's X-ray microscopy facilities, including specimen advice, microscopy and data reconstruction and to provide local and national leadership in the data management and image analysis requirements of the centre.

In this role you will:

. coordinate and deliver the training program for user groups in the X-ray and image analysis sections
. instruct, train and assist users in their research and develop a user portfolio that demonstrates a quality user experience and effective user outcomes in the development and application of X-ray and image analysis techniques
. work with colleagues to facilitate microscopy experiments for users, and provide laboratory and microscopy services to commercial clients of the centre.
. develop and modify procedures and operational techniques to achieve continuous improvement
. monitor the X-ray facility budget and work closely with the Laboratory Manager to procure and advise on X-ray/image analysis instrumentation.

To qualify for this role, you will demonstrate:

. experience in coordinating the day-today running of microscopy instrumentation, including considering in standard specimen preparation techniques for microscopy
. knowledge, skills and experience in the management of data including the transfer, archiving and disposal of data, and the proper aggregation of metadata
. in-depth, specialist knowledge and skills in the 3D data segmentation, image rendering and image analysis by means of advanced image analysis software, including an ability to write computer codes that execute advanced analyses on various forms of microscopy data
. an ability to ascertain ongoing microscopy and microanalysis trends, methodological developments and the needs of users across the X-ray area.

All applications must be submitted via the University of Sydney careers website.� Visit sydney.edu.au/positions and search by the reference number for more information and to apply.

CLOSING DATE: 10 April 2012, 1pm

The University is an Equal Opportunity employer committed to equity, diversity and social inclusion. Applications from equity target groups and women are encouraged.

� The University of Sydney

==============================Original Headers==============================
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16, 37 -- Subject: JOB OPPORTUNITY: PROFESSIONAL OFFICER - X-RAY MICROSCOPY AT THE
16, 37 -- UNIVERSITY OF SYDNEY
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PROFESSIONAL OFFICER - LIGHT AND OPTICAL MICROSCOPIST
SYDNEY MICROSCOPY & MICROANALYSIS
THE UNIVERSITY OF SYDNEY
REFERENCE NO. 372/0312

. Coordinate light/optical facilities in consultation with the Laboratory Manager
. Facilitate microscopy experiments for users, and provide laboratory and microscopy services to commercial clients of the centre
. Full-time continuing, remuneration package: $92K p.a. which includes leave loading and up to 17% super

Sydney Microscopy & Microanalysis (SMM) aims to be a world-leading facility for modern microscopy and microanalysis, offering premier instruments, services and training to researchers from across Sydney and around Australia, and providing strong leadership in the national and international characterisation communities.

Reporting to the Laboratory Manager, you will coordinate the centre's light and optical facilities, focussing primarily on the needs of users in, microscopy, microanalysis, and analysis and interpretation of images and data, so that these facilities provide effective, high-quality outcomes for users.

In this role you will:

. instruct, demonstrate, train and assist users in their research and develop a user portfolio that demonstrates a quality user experience and effective user outcomes in the development and application of light/optical-based techniques
. work with colleagues to facilitate microscopy experiments for users, and provide laboratory and microscopy services to commercial clients of the Centre
. develop and modify procedures and operational techniques to achieve continuous improvement
. report on use of the light/optical facility, ensuring instrument logs are completed accurately and user information is reported.

To qualify for this role, you will demonstrate:

. extensive experience and expertise in light-optical microscopy and associated specimen preparation techniques and data analysis including know-how of illumination modes and contrast enhancement methods, confocal and fluorescence microscopy techniques and immuno-labelling
. skills in advanced light/optical techniques, including, FRET, FLIM, FRAP, ratiometric and live-cell, along with the associated analysis techniques
. experience in devising microscopy solutions for research including post-processing and image analysis for the transformation of data into publishable information and results
. advanced problem solving skills and demonstrated ability to run and maintain sophisticated scientific instrumentation

All applications must be submitted via the University of Sydney careers website.� Visit sydney.edu.au/positions and search by the reference number for more information and to apply.

CLOSING DATE: 10 April 2012, 1pm

The University is an Equal Opportunity employer committed to equity, diversity and social inclusion. Applications from equity target groups and women are encouraged.

� The University of Sydney

==============================Original Headers==============================
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14, 31 -- To: "Microscopy-at-Microscopy.Com" {Microscopy-at-Microscopy.Com}
14, 31 -- Subject: JOB OPPORTUNITY: PROFESSIONAL OFFICER - LIGHT AND OPTICAL
14, 31 -- MICROSCOPIST AT THE UNIVERSITY OF SYDNEY
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14, 31 -- MICROSCOPIST AT THE UNIVERSITY OF SYDNEY
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I need help from someone familiar with the DIC (Nomarski) equipment used
with the Nikon Microphot series of microscopes. I am interested in
transmitted light DIC with 160 mm objectives, not epi-illumination DIC.
My understanding is that you need the Universal condenser, a DIC slider,
and either a dedicated DIC intermediate nosepiece mount, or the EPI-FL3
unit with the 1.25x intermediate nosepiece mount, which has a slot for
the slider.
I purchased a condenser and Nikon slider (#59234) that was supposed to
be for the Microphot. When used with my EPI-FL3 unit, the slider has
the proper width and thickness to fit into the intermediate nosepiece
mount, but it is much too long. It runs into the nosepiece before even
the blank hole in the slider is in proper position. The slider measures
125 x 38 x 15mm. I think it would need to be about 40mm shorter to
operate properly with the EPI-FL3 and the standard nosepieces I have.

What are the dimensions of the proper slider that goes with the
EPI-FL3? Is there a special nosepiece that will allow my slider to
operate with the EPI-FL3? What configuration is my slider intended for?

I would also be interested in hearing from anyone interested in selling
the proper DIC components to operate with my scopes (Microphot FXA and SA).

Ralph Common

==============================Original Headers==============================
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Hello Listers,

One of our Pathologists wants to know which fixative and buffer
combination are the best for the preservation of glycogen at the EM
level.

We are getting more and more muscle biopsies with glycogen storage
issues and she wants to make sure we are fixing them the best way
possible.

Currently we use a 4% glutaraldehdye in 0.1 M sodium cacodylate buffer.

Any and all suggestions are gratefully accepted.

May the best glycogen fixative win!

Thanks in advance,

Paula :-)

--
Paula Sicurello, HTL (ASCP)
Supervisor, Clinical Electron Microscopy Laboratory
Duke University Health System
Rm.#251M, Duke South, Green Zone
Durham, North Carolina 27710
P: �919.684.2091

==============================Original Headers==============================
10, 29 -- From patpxs-at-gmail.com Mon Apr 2 12:51:35 2012
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10, 29 -- Subject: Glycogen Preservation-best fixative?
10, 29 -- From: Paula Sicurello {patpxs-at-gmail.com}
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Hi Paula,

Wim de Bruijn has done a lot of work for the specific staining of glycogen, using OsO4 and Potassium Ferro/Ferricyanide mixtures. I do not have the literature available, but a search on Google will give you plenty of leads.

Good luck,

Jan

Jan Leunissen PhD
Aurion

On 3/04/2012, at 5:51 AM, patpxs-at-gmail.com wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hello Listers,
}
} One of our Pathologists wants to know which fixative and buffer
} combination are the best for the preservation of glycogen at the EM
} level.
}
} We are getting more and more muscle biopsies with glycogen storage
} issues and she wants to make sure we are fixing them the best way
} possible.
}
} Currently we use a 4% glutaraldehdye in 0.1 M sodium cacodylate buffer.
}
} Any and all suggestions are gratefully accepted.
}
} May the best glycogen fixative win!
}
} Thanks in advance,
}
} Paula :-)
}
} --
} Paula Sicurello, HTL (ASCP)
} Supervisor, Clinical Electron Microscopy Laboratory
} Duke University Health System
} Rm.#251M, Duke South, Green Zone
} Durham, North Carolina 27710
} P: 919.684.2091
}
}
} ==============================Original Headers==============================
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The Spring MSORV meeting (Microscopy Society of the Ohio River Valley)
will be held on Wednesday, April 18th at UES, Inc., 4401 Dayton-Xenia
Rd., Dayton, OH 45432 3-7:00 PM.

An agenda is posted on the MSORV website, www.msorv.org. We are very
excited to have one of MSA's Past Presidents, Dr. Dave Piston, and MAS's
Past President, Dr. John Henry Scott, as our keynote speakers. Speaker
abstracts are available on the MSORV website, along with an agenda, maps
and directions.

To attend, you will have to pre-register in order for us to provide the
caterer with a head count and to provide a list of names for the
Receptionist at UES. There is a $15 registration fee to attend for
professional non-members and guests and $5 for student non-members. The
registration fee will be waived if you choose to join at the meeting.
Registration fees will be collected at the door.
MSORV members (student and professional) do not have to pay a
registration fee, but everyone planning to attend must pre-register!

Please send an e-mail to Pam Lloyd, MSORV's Secretary, at
pamela.lloyd.ctr-at-wpafb.af.mil by April 11, 2012 to confirm your
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I have always had good success with 4% glutaraldehyde, used with
phosphate buffer. As far as I know, cacodylate should also be OK. For
preservation of glycogen, it is essential that en-bloc UA is NOT used.
It dissolves the glycogen, or if there is a lot of glycogen, causes it
to form gummy clumps. I follow fixation with 3 hours in 1% OsO4, and
normal processing (alcohol dehydration, infiltration with "Epon"
substitute resin in propylene oxide).

It is also important that the samples be fixed promptly, and not set in
buffer for a prolonged period before processing.

Ralph Common

==============================Original Headers==============================
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Hi Paula,

About three years ago in late March, 2009 there was a discussion
about glycogen preservation on this listserver so you might want to
search the archives for that information. Here are two references
relevant to that discussion:

1) K.K. Rybicka, 1996. Tissue & Cell 28 (3) 253-267.
2) Microscopy Today, October 1994. "Glycogen Granules Revisited".

Also, here is a recipe that proved useful for us at the Dana-Farber
Cancer Institute in preserving glycogen rosettes in liver biopsies of
pediatric cancer patients. Tissue was post-fixed with osmium potassium
ferrocyanide after glut fix followed by alcohol and propylene oxide
dehydration and epon embedment. UA enbloc staining is not recommended.

2 mls. 2% aqueous osmium
2 mls. buffer (ie. 0.1M Na-phosphate, pH 7.4)
0.06 gms potassium ferrocyanide

The OPF solution is prepared with thorough mixing just prior to
use and applied to tissue for 2 hours in refrigerator.

Best of luck, Don

Donald Gantz
Dept. Physiology & Biophysics
Boston Univ. School of Medicine

On 4/2/2012 1:52 PM, patpxs-at-gmail.com wrote:
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} ----------------------------------------------------------------------------
}
} Hello Listers,
}
} One of our Pathologists wants to know which fixative and buffer
} combination are the best for the preservation of glycogen at the EM
} level.
}
} We are getting more and more muscle biopsies with glycogen storage
} issues and she wants to make sure we are fixing them the best way
} possible.
}
} Currently we use a 4% glutaraldehdye in 0.1 M sodium cacodylate buffer.
}
} Any and all suggestions are gratefully accepted.
}
} May the best glycogen fixative win!
}
} Thanks in advance,
}
} Paula :-)
}

==============================Original Headers==============================
10, 24 -- From gantz-at-bu.edu Mon Apr 2 13:39:40 2012
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Dear Paula,

You need to use reduced osmium tetroxide (1% OsO4 + 1.5% K4Fe(CN)6 in 0.1M caclodylate buffer) instead osmium tetroxide for postfixation. You will get beautiful electron dense glycogen granules. Uranyl acetate en block will not dissolve them but will make them less contrasty. Primary fixation is not that important in this case as well.

Sincerely,
Alex
--
Dr. Aleksandr Mironov MD, PhD
Senior Experimental Officer
D.1527, M.Smith Building
EM Core Facility, Faculty of Life Sciences
University of Manchester
Oxford Road
Manchester
M13 9PT
UK

Tel. +44-(0)161-275-5645
Fax. +44-(0)161-275-5171
E-mail: Aleksandr.Mironov-at-manchester.ac.uk
http://www.ls.manchester.ac.uk/research/facilities/electronmicroscopy/

________________________________________
X-from: patpxs-at-gmail.com [patpxs-at-gmail.com]
Sent: 02 April 2012 18:56
To: Aleksandr Mironov

Hello Listers,

One of our Pathologists wants to know which fixative and buffer
combination are the best for the preservation of glycogen at the EM
level.

We are getting more and more muscle biopsies with glycogen storage
issues and she wants to make sure we are fixing them the best way
possible.

Currently we use a 4% glutaraldehdye in 0.1 M sodium cacodylate buffer.

Any and all suggestions are gratefully accepted.

May the best glycogen fixative win!

Thanks in advance,

Paula :-)

--
Paula Sicurello, HTL (ASCP)
Supervisor, Clinical Electron Microscopy Laboratory
Duke University Health System
Rm.#251M, Duke South, Green Zone
Durham, North Carolina 27710
P: 919.684.2091

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Hello Listers,

Based on the responses I'm getting, I know I wasn't clear enough with
my question. So I will rephrase it.

What is the best primary fixative?

or put another way: What buffer is the best for preserving glycogen,
used with what fixative?

Our processing protocol includes the osmium step, we DO NOT en-bloc
with UA. [On an aside, is en-bloc UA really all that beneficial?
Past results in my experience is NO.]

Thanks in advance, again!

Paula :-)

--
Paula Sicurello, HTL (ASCP)
Supervisor, Clinical Electron Microscopy Laboratory
Duke University Health System
Rm.#251M, Duke South, Green Zone
Durham, North Carolina 27710
P: �919.684.2091

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9, 29 -- Subject: Best Fix for glycogen-clarification
9, 29 -- From: Paula Sicurello {patpxs-at-gmail.com}
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Dear Paula,

To my experience it does not matter much which primary fixative and buffer you will use - glutaraldehyde, formaldehyde, cacodylate, PBS, phosphate, zwitterionic buffers. The most important step is the usage of reduced osmium (instead of standard osmium tetroxide) - then your glycogen will be impossible to miss.

Regards,
Alex
--
Dr. Aleksandr Mironov MD, PhD
Senior Experimental Officer
D.1527, M.Smith Building
EM Core Facility, Faculty of Life Sciences
University of Manchester
Oxford Road
Manchester
M13 9PT
UK

Tel. +44-(0)161-275-5645
Fax. +44-(0)161-275-5171
E-mail: Aleksandr.Mironov-at-manchester.ac.uk
http://www.ls.manchester.ac.uk/research/facilities/electronmicroscopy/

________________________________________
X-from: patpxs-at-gmail.com [patpxs-at-gmail.com]
Sent: 02 April 2012 21:54
To: Aleksandr Mironov

Hello Listers,

Based on the responses I'm getting, I know I wasn't clear enough with
my question. So I will rephrase it.

What is the best primary fixative?

or put another way: What buffer is the best for preserving glycogen,
used with what fixative?

Our processing protocol includes the osmium step, we DO NOT en-bloc
with UA. [On an aside, is en-bloc UA really all that beneficial?
Past results in my experience is NO.]

Thanks in advance, again!

Paula :-)

--
Paula Sicurello, HTL (ASCP)
Supervisor, Clinical Electron Microscopy Laboratory
Duke University Health System
Rm.#251M, Duke South, Green Zone
Durham, North Carolina 27710
P: 919.684.2091

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Email: bob.price-at-uscmed.sc.edu Name: Bob Price

Organization: Univ South Carolina

Title-Subject: [Filtered] Workshop on Basic Confocal Microscopy

Message: The University of South Carolina School of Medicine Instrumentation Resource Facility and
the Clemson University Microscopy Facilities will be hosting the 8th Annual Workshop on Basic
Confocal Microscopy at Clemson University from June 11th through the 15th

The workshop is directed towards beginning and intermediate users of confocal microscopes and
involves a series of lectures (specimen preparation, labeling strategies, proper set-up of
instrument operating parameters, proper handling of 2D and 3D confocal images in programs such as
Adobe Photoshop and AMIRA), hands on specimen preparation, and time on a number of different point
scanning and spinning disk confocal microscopes.

Companies scheduled to have instruments and applications experts on-site include Leica, Nikon,
Olympus, Perkin Elmer, Zeiss and Intelligent Imaging Innovations (3i).

Faculty will include Dr. Jay Jerome of Vanderbilt University, Dr. Ralph Albrecht of the University
of Wisconsin Madison, Dr. John Mackenzie of North Carolina State University, Dr. Tom Trusk of the
Medical University of South Carolina, and Dr. Bob Price of the University of South Carolina.

For further information please see: http://dba.med.sc.edu/price/irf/irf.htm or contact Anna McNeal
(anna.mcneal-at-uscmed.sc.edu {mailto:anna.mcneal-at-uscmed.sc.edu} ), Bob Price
(bob.price-at-uscmed.sc.edu {mailto:bob.price-at-uscmed.sc.edu} ) or JoAn Hudson
(joanh-at-clemson.edu {mailto:joanh-at-clemson.edu} )

Bob Price
Research Professor
USC School of Medicine
6439 Garner's Ferry Road
Columbia, SC 29209
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Hi Owen,

We had recently an extensive discussion in the List about the preparation of HIV particles for TEM.
The actual question is not whether or not the virus is inactivated and still dangerous after fixation.
The real issue is regulations. BSL-2 implicates that specific measures�are undertaken not only to protect the person who manipulates the samples, but also all his colleagues who are perhaps not even aware that someone is working with a BSL-2 virus next bench/door. Please read carefully the regulations about BSL-2 and, as already advised, discuss with the right persons about it. I mean, not electron microscopists but security/health managers. There are strict regulations and protocols which must be strictly followed, it is not as if anyone can do anything with BSL samples.

Best regards,
Stephane

�

----- Original Message -----
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Cc:
Sent: Friday, March 30, 2012 4:49 PM

All,

A scientist has asked to image porcine virus in our FESEM.� She says that it is listed as a biosafety level 2, but has been disinfected to } 99% level with 10% glutaraldehyde.� Are any of you working with this virus?� I'm not familiar with the safety protocols.� Is this a safe project for a core facility?�

Thanks in advance.

Owen

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Good morning,

We are having issues with wrinkles in our 1 to 4-micron resin sections. Does anyone have a recommendation for adding acetone (percentage?) to the flotation water in order to reduce the problem?

We're using an Embed812-Araldite6005 mixture and, due to specimen size, block face is approximately 5mm high x 3mm wide. We place drops of de-ionized water on clean slides, then position one section on each drop. The sections are allowed to float at room temperature for a bit to relax on their own and then the slides are moved to hotplates--first at 80 degrees Celsius for approximately 5-10 seconds and then at 60 degrees Celsius until completely dry (15 minutes minimum).

Thank you for your time,

Jaclynn Lett

Senior Research Technician, EM Facility
Research Center for Auditory and Vestibular Studies
Department of Otolaryngology
Washington University School of Medicine
660 S Euclid Ave, Campus Box 8115
St. Louis, MO 63110

Office: 314-747-7257
Fax: 314-747-7230
Email: lettj-at-ent.wustl.edu
http://otocore.wustl.edu

The materials in this email are private and may contain Protected Health Information. If you are not the intended recipient, be advised that any unauthorized use, disclosure, copying, distribution or the taking of any action in reliance on the contents of this information is strictly prohibited. If you have received this email in error, please immediately notify the sender via telephone or return email.

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Dear Jaclynn,

not easy to solve, I guess...
wrinkling and "curving/screwing up" sections with usual thickness (1-2 �m) perhaps IMHO depend on knife type (glass or diamond), sectioning parameters (like shape of trimmed faces, section velocity, chosen thickness) , naturally on block dimensions and last but not least on properties of resin.

Since Embed812(Epon812)-Araldite has been reported to be sometimes brittle (also depending on the temperature and time modes of polymerization), it would be also of interest which hardness/softness you chose in your resin mixture... Such big areas (5 x 3 mm) perhaps might be sectioned better when really soft (depending too on your object and what you are doing / plan to do after sectioning, I guess).
4 �m thickness really is a challenge (I have done serial 2�m resin block sectioning of rat brain - whole hypothalamus region including ventricle region - 3-3.5 x 4 mm in my old times) after a while without problems (Epon 812 at that time, soooft mixture).... might function also with liver... other tissues, I guess, might be really problematic.
Acetone in the knife boat (obsolete for the knife), the flotation water or Chloroform vapors (CAVE: not very healthy) might be of no real advantage here, prolonged drying in a drop of water on the slide (under an ancient light bulb) perhaps could help a little bit...
Unfortunately my knowledge (and "wisdom") now is at its end, sorry for that...
best wishes and good luck, hoping another lister HAS a real solution....

Happy Easter,
cordially yours
Wolfgang MUSS PhD
Salzburg, Austria

} -----Urspr�ngliche Nachricht-----
} Von: LettJ-at-ent.wustl.edu [mailto:LettJ-at-ent.wustl.edu]
} Gesendet: Dienstag, 03. April 2012 18:01
} An: Mu� Wolfgang
} Betreff: [Microscopy] wrinkled thick sections
}
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} Good morning,
}
} We are having issues with wrinkles in our 1 to 4-micron resin sections.
} Does anyone have a recommendation for adding acetone (percentage?) to
} the flotation water in order to reduce the problem?
}
} We're using an Embed812-Araldite6005 mixture and, due to specimen size,
} block face is approximately 5mm high x 3mm wide.
}
} We place drops of de-ionized water on clean slides, then position one
} section on each drop.
} The sections are allowed to float at room temperature for a bit to
} relax on their own and then the slides are moved to hotplates -- first
} at 80 degrees Celsius for approximately 5-10 seconds and then at 60
} degrees Celsius until completely dry (15 minutes minimum).
}
}
} Thank you for your time,
}
} Jaclynn Lett
}
} Senior Research Technician, EM Facility
} Research Center for Auditory and Vestibular Studies
} Department of Otolaryngology
} Washington University School of Medicine
} 660 S Euclid Ave, Campus Box 8115
} St. Louis, MO 63110
}
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7, 39 -- From W.Muss-at-salk.at Tue Apr 3 12:35:13 2012
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I'm not familiar with the Microphot in particular, but I can tell you that
vendor support for older microscopes, especially fixed tube length
systems, is pretty limited. It's a bit like vintage camera equipment - you
can get beautiful images from it, but replacing a broken part or upgrading
means being able to obtain vintage equipment from sources such as eBay,
and generally being plugged in to a specialized group of legacy equipment
users and hobbyists who will have the information you need about your
older system.

You didn't make clear who you got the slider from. Was it an eBay sale or
from Nikon itself? In any event, I recommend trying to find Nikon
literature from that era so that you can have all the part numbers for the
Microphot compatible components you might need.

Science-Info.net is a good resource. Here is it's list of older Nikon
manuals, albeit, I don't see any specifically for the Microphot:

http://www.science-info.net/docs/Nikon/

There's a dealer going by the name Classic Optics that might be able to
help you with literature and possibly even the part itself:

http://www.science-info.net/docs/Nikon/

I also *highly* recommend the Microscope Yahoo Group, which is a fairly
high-traffic list with many members who are highly knowledgeable about
older microscopes:

http://tech.dir.groups.yahoo.com/group/Microscope/

All the best,
Peter G. Werner
Program Assistant, Merritt College Microscopy Program
President, San Francisco Microscopical Society

} I need help from someone familiar with the DIC (Nomarski) equipment used
} with the Nikon Microphot series of microscopes. I am interested in
} transmitted light DIC with 160 mm objectives, not epi-illumination DIC.
} My understanding is that you need the Universal condenser, a DIC slider,
} and either a dedicated DIC intermediate nosepiece mount, or the EPI-FL3
} unit with the 1.25x intermediate nosepiece mount, which has a slot for
} the slider.
} I purchased a condenser and Nikon slider (#59234) that was supposed to
} be for the Microphot. When used with my EPI-FL3 unit, the slider has
} the proper width and thickness to fit into the intermediate nosepiece
} mount, but it is much too long. It runs into the nosepiece before even
} the blank hole in the slider is in proper position. The slider measures
} 125 x 38 x 15mm. I think it would need to be about 40mm shorter to
} operate properly with the EPI-FL3 and the standard nosepieces I have.
}
} What are the dimensions of the proper slider that goes with the
} EPI-FL3? Is there a special nosepiece that will allow my slider to
} operate with the EPI-FL3? What configuration is my slider intended for?
}
} I would also be interested in hearing from anyone interested in selling
} the proper DIC components to operate with my scopes (Microphot FXA and
} SA).
}
} Ralph Common

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Marissa;
CMP means Chemical Mechanical Polish. That means that the silicon is lightly etched while it is being polished. Any amorphization is so little that it could not be detected by TEM. it is almost ready for growth of gate oxide at this point. The epitaxial layer grown with silane is usually not pure silane, but accompanied by some controlled impurities to dope the surface of the wafer. You will never see any evidence in a TEM of the interface between the epitaxial layer and the underlying wafer in semiconductor quality silicon. The only thing you may see at the surface of either CMP or epitaxial silicon would be a thin layer of oxide, called native oxide. The thickness can depend on how long the silicon was exposed to air and what other species may be adsorbed onto the surface. The native oxide would generally be only a few angstroms thick.

John Mardinly, ASU

On Apr 2, 2012, at 5:18 PM, "microscopylistserver-noreply-at-microscopy.com" {microscopylistserver-noreply-at-microscopy.com} wrote:

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} Title-Subject: [Filtered] Amorphous damage from CMP wafer preparation
}
} Message: During the Si wafer growth and preparation process, how much damage is produced from the
} CMP process? Also, I've read that an epitaxial layer is grown on the wafer by passing silane gas
} over the surface. Will someone comment on the way these processes impact the structure of the Si at
} the surface of the wafer?
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} Marissa
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On Apr 3, 2012, at 12:49 PM, John.Mardinly-at-asu.edu wrote:

} Marissa;
} CMP means Chemical Mechanical Polish. That means that the
} silicon is lightly etched while it is being polished. Any
} amorphization is so little that it could not be detected by TEM. it
} is almost ready for growth of gate oxide at this point. The
} epitaxial layer grown with silane is usually not pure silane, but
} accompanied by some controlled impurities to dope the surface of the
} wafer. You will never see any evidence in a TEM of the interface
} between the epitaxial layer and the underlying wafer in
} semiconductor quality silicon. The only thing you may see at the
} surface of either CMP or epitaxial silicon would be a thin layer of
} oxide, called native oxide. The thickness can depend on how long the
} silicon was exposed to air and what other species may be adsorbed
} onto the surface. The native oxide would generally be only a few
} angstroms thick.
}
} John Mardinly, ASU
}
} } Email: mlibbee-at-gmail.com Name: Marissa
} }
} } Organization: LBL- NCEM
} }
} } Title-Subject: [Filtered] Amorphous damage from CMP wafer preparation
} }
} } Message: During the Si wafer growth and preparation process, how
} } much damage is produced from the
} } CMP process? Also, I've read that an epitaxial layer is grown on
} } the wafer by passing silane gas
} } over the surface. Will someone comment on the way these processes
} } impact the structure of the Si at
} } the surface of the wafer?
} }
} } Much obliged,
} } Marissa
} }
Hi John and Marissa,
Could grazing-incidence electron diffraction, used, for example, by
Larry Marks (sp?), detect and/or quantitate surface changes produced
by CMP and silane?
Yours,
Bill

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Hello Jaclynn,
I cut 1u sections (and 5u sections) with Diatome diamond knives mostly samples embedded in Polybed 812. I would transfer them onto a glass slide with a wooden stick or a loop. I would put the slide on a hotplate 60C and then invert a large glass Petri dish over the slide. I would place a cotton swab dipped in acetone under the dish with the slide. The acetone vapor + the heat would flatten the sections and as the drop of water evaporated the sections would anneal to the slide. It worked great for me. Good luck
Dean Abel
Biological Sciences
University of Iowa
Iowa City IA USA

We are having issues with wrinkles in our 1 to 4-micron resin sections. Does anyone have a recommendation for adding acetone (percentage?) to the flotation water in order to reduce the problem? We're using an Embed812-Araldite6005 mixture and, due to specimen size, block face is approximately 5mm high x 3mm wide. We place drops of de-ionized water on clean slides, then position one section on each drop. The sections are allowed to float at room temperature for a bit to relax on their own and then the slides are moved to hotplates--first at 80 degrees Celsius for approximately 5-10 seconds and then at 60 degrees Celsius until completely dry (15 minutes minimum). Thank you for your time, Jaclynn Lett

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Dear listeners

I know only one system to clean samples inside a SEM to avoid
contamination dammage. Would you please inform if you know an alternate
choice to EVACTRON from XEI scientific ?
Thanks a lot.

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

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} Date: Tue, 3 Apr 2012 13:54:15 -0700
} From: {AssociationManagement-at-microscopy.org}
} Reply-To: Mauricio Bravo {mauricio.bravo-at-maxim-ic.com}
} realname - Mauricio Bravo
} Email - mauricio.bravo-at-maxim-ic.com
} ORGANIZATION - Maxim Integrated Products
} EDUCATION - Graduate College
} LOCATION - Dallas, Texas
} SUBJECT_OF_QUESTION - FIB circuit edits on microchips using copper
} metallization
} QUESTION - I am a FIB operator, I operate a FEI Altura 855 DualBeam
} machine. Normally we use gallium ions and Iodine to selectively cut
} aluminum metal lines without damaging the insulator around it; now I
} need to prepare to receive ICs built using copper metal lines. I
} contacted FEI and they suggest to use water as a chemical precursor
} to cleanly cut the copper without much damage to metal barriers
} protecting insulators. My question is, are there other options to
} copper milling using water? If so, where I can find literature
} explaining the other options? Thanks very much for your time.
}
} Mauricio Bravo

--

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There is an inherent difference in GAE for FIB milling Al vs. Cu
conductors. In case of Al you are using precursor (commonly I, Br, or
Cl2 being the best if you are lucky to have it) to volatilize metal and
remove it from the repair area. In case of Cu there is no know (at least
to me) methods to volatilize the metal in FIB, so precursor is used to
inhibit FIB damage to surrounding/underlying dielectrics and (hopefully)
to oxidize the Cu to non-conductive state. H2O is a known inhibitor of
Ga+ FIB etching for typical dielectric materials: SiO2, SiN, and even a
CDO, but in principle you can find plenty of other (organic) precursors
with stronger absorbance to dielectric then to Cu and thus providing
etch inhibition. Some of such precursors are mildly corrosive and may
also aid in oxidizing Cu.

With properly designed GAE recipes and procedures you can get very good
results using H2O precursor for etching Cu conductors over SiO2 and SiN
dielectrics (I am sure it would also work for CDO, but I have not tried):

http://partbeamsystech.academia.edu/RayValery/Papers/520167/Advanced_FIB_Circuit_Edit_Cu_Line_Deprocessing

Just remember that standard GAE recipes provided by OEM of your FIB tool
may be suboptimal and need some serious tweaking to get best results for
etching extremely thin and/or very thick Cu lines.

However etching Cu conductors with H2O precursor would not work if you
have organic Low-K dielectric around or immediatelly below your Cu line,
as H2O will enhance etching of such dielectric layers instead of
inhibiting it. In this case your best bet is to contact Vladimir Makarov
and Leo Arbatman of Tiza Labs: http://tiza-lab.com/ (no any interest
here - just the only known to me commercial source of FIB GAE technology
for Cu etching over organics).

Good luck :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 4/4/2012 9:25 AM, oshel1pe-at-cmich.edu wrote:
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} as well as to the list.
} Using the "reply" function in your email does *not* send your answer
} to the person asking the question.
} Please copy their email address from their question.
} ****************************************************************************************
} } Date: Tue, 3 Apr 2012 13:54:15 -0700
} } From: {AssociationManagement-at-microscopy.org}
} } Reply-To: Mauricio Bravo {mauricio.bravo-at-maxim-ic.com}
} } realname - Mauricio Bravo
} } Email - mauricio.bravo-at-maxim-ic.com
} } ORGANIZATION - Maxim Integrated Products
} } EDUCATION - Graduate College
} } LOCATION - Dallas, Texas
} } SUBJECT_OF_QUESTION - FIB circuit edits on microchips using copper
} } metallization
} } QUESTION - I am a FIB operator, I operate a FEI Altura 855 DualBeam
} } machine. Normally we use gallium ions and Iodine to selectively cut
} } aluminum metal lines without damaging the insulator around it; now I
} } need to prepare to receive ICs built using copper metal lines. I
} } contacted FEI and they suggest to use water as a chemical precursor
} } to cleanly cut the copper without much damage to metal barriers
} } protecting insulators. My question is, are there other options to
} } copper milling using water? If so, where I can find literature
} } explaining the other options? Thanks very much for your time.
} }
} } Mauricio Bravo
}

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Bill, At that point it becomes a research project.

John Mardinly, ASU

On Apr 3, 2012, at 5:30 PM, "Bill & Sue Tivol" {wtivol-at-sbcglobal.net} wrote:

}
} On Apr 3, 2012, at 12:49 PM, John.Mardinly-at-asu.edu wrote:
}
} } Marissa;
} } CMP means Chemical Mechanical Polish. That means that the silicon is lightly etched while it is being polished. Any amorphization is so little that it could not be detected by TEM. it is almost ready for growth of gate oxide at this point. The epitaxial layer grown with silane is usually not pure silane, but accompanied by some controlled impurities to dope the surface of the wafer. You will never see any evidence in a TEM of the interface between the epitaxial layer and the underlying wafer in semiconductor quality silicon. The only thing you may see at the surface of either CMP or epitaxial silicon would be a thin layer of oxide, called native oxide. The thickness can depend on how long the silicon was exposed to air and what other species may be adsorbed onto the surface. The native oxide would generally be only a few angstroms thick.
} }
} } John Mardinly, ASU
} }
} } } Email: mlibbee-at-gmail.com Name: Marissa
} } }
} } } Organization: LBL- NCEM
} } }
} } } Title-Subject: [Filtered] Amorphous damage from CMP wafer preparation
} } }
} } } Message: During the Si wafer growth and preparation process, how much damage is produced from the
} } } CMP process? Also, I've read that an epitaxial layer is grown on the wafer by passing silane gas
} } } over the surface. Will someone comment on the way these processes impact the structure of the Si at
} } } the surface of the wafer?
} } }
} } } Much obliged,
} } } Marissa
} } }
} Hi John and Marissa,
} Could grazing-incidence electron diffraction, used, for example, by Larry Marks (sp?), detect and/or quantitate surface changes produced by CMP and silane?
} Yours,
} Bill
}
}
}

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Carl Zeiss has an option for cleaning and avoiding contamination in an SEM.
They bring a gas injection needle in close to the sample and allow oxygen in
through the system around the sample. In the presence of the electron beam,
activated species are created which will clean the sample in much the same
way as a plasma cleaner or a UVO-Cleaning system would clean the sample. I
suggest that you look at the brochure. They even show the removal of
previously contaminated area.

Since being introduced to this system by reading their brochure, I've
thought about alternatives to it. Could you put an oxygen leak into the SEM
and obtain the same result? I've made a few assumptions and did a few
vacuum calculations to see whether it is feasible. First, the Zeiss gas
injection system can give you a local higher concentration of gas at the
sample with an overall lower background pressure, i.e. the thoughput (the
gas load) is smaller. This is good since oxygen would be deleterious for
the electron gun. But what about if you simply put the leak into the SEM
chamber. I figure that for reasonable cleaning action, that you would need
about a pressure of 1E-6 Torr at the sample. At this pressure, a monolayer
of gas arrives at the surface every second. I don't know what the cross
section is for the oxygen molecule for activation or ionization, but I'm
sure that it is less than one. The sticking coefficient of oxygen on the
surface will be dependent on the material, but it also is less than one.
What this means is that for a contaminated sample, you would probably need
to scan the beam over the sample at a lower magnification for several
minutes prior to going to a higher magnification. For a sample that is
relatively clean and you just want to maintain cleaning, you could probably
just start imaging right away.

I've done the calculation for the gas flow needed making the assumption that
the effective pump speed for a high vacuum pump at the SEM chamber would be
about 200 l/s. This is probably a conservative value for most SEMs with a
turbopump placed right on the chamber. Using the expression Q=SP where Q is
the throughput (Torr-l/sec), S is the pump speed (l/sec) and P is the
pressure (Torr), at 1E-6 Torr and a pump speed of 200 l/sec, the throughput
would be 2E-4Torr-l/sec. This converts to 0.016 sccm. This is a very small
gas leak that is easily achievable with a moderately priced leak valve. I
don't know what the effect of prolonged use of the oxygen leak into the
chamber would be on the various types of electron sources. The gun areas
are typically differentially pumped relative to the chamber so that this is
good. Also, the ion pumps used on LaB6 and field emission sources pump
oxygen very effectively. I suspect that this level of oxygen in a system
would not significantly change the lifetime of a filament in most high
vacuum SEM systems.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: Nicolas.Stephant-at-univ-nantes.fr
[mailto:Nicolas.Stephant-at-univ-nantes.fr]
Sent: Wednesday, April 04, 2012 1:23 AM
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Dear listeners

I know only one system to clean samples inside a SEM to avoid contamination
dammage. Would you please inform if you know an alternate choice to EVACTRON
from XEI scientific ?
Thanks a lot.

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en produire
une image"

C.G Jung

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Email: leswes-at-shaw.ca Name: Lesley Weston

Organization: UBC

Title-Subject: [Filtered] Re Wrinkled thick sections

Message: You might find that your resin could be adjusted. I used to cut very-large-area, very thick
sections by embedding in a harder mix than Epon/Araldite, using Jembed with NMA and DDSA, but then
my tissue was pretty hard too. I handled the sections as you describe, but one difference was that I
cut the sections dry on a glass knife, using two pairs of fine forceps to pick them up off the knife
face and transfer them to the water drops on the slides. I was able to cut serials of blocks 5 mm
square at 1 or 2 mu like this, with no wrinkling. For reorientating smaller pieces of the block for
thin sectioning, I would cut a few sections at 10 mu or more. If your tissue is soft so that you
need to use the softer resin, you could still try cutting the sections dry.

Lesly Weston.

"Good morning,

We are having issues with wrinkles in our 1 to 4-micron resin sections. Does anyone have a
recommendation for adding acetone (percentage?) to the flotation water in order to reduce the problem?

We're using an Embed812-Araldite6005 mixture and, due to specimen size, block face is approximately
5mm high x 3mm wide. We place drops of de-ionized water on clean slides, then position one section
on each drop. The sections are allowed to float at room temperature for a bit to relax on their own
and then the slides are moved to hotplates--first at 80 degrees Celsius for approximately 5-10
seconds and then at 60 degrees Celsius until completely dry (15 minutes minimum).

Thank you for your time,

Jaclynn Lett"

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Marissa asked about characterizing surface changes on Silicon wafers as a
result of CMP and epitaxial deposition.
John Mardinly and Bill Tivol commented regarding the general characteristics
of the materials, especially the ultrathin films involved, and briefly
discussed TEM and grazing incidence electron diffraction.
I have had great success in looking at Silicon surfaces with the AFM (atomic
force microscope). It is exquisitely sensitive to small height changes,
down to the atomic scale. An example AFM image of an epitaxial layer is
shown at http://www.asmicro.com/corporate/yale.htm.
AFM is the tool of choice in the semiconductor industry for characterizing
surface changes. It is easy to see scratches and to measure surface
roughness down to rms roughness { 0.1 nm.

Commercial disclaimer: My company provides AFM analytical services and
also buys, refurbishes, sells and installs used (second-hand) AFMs. For
further information, please contact me offline.
regards,
Don
=============================================
Don Chernoff, Ph.D., President
Advanced Surface Microscopy, Inc. E-Mail: donc-at-asmicro.com
3250 N. Post Rd., Ste. 120 Voice: 317-895-5630
INDIANAPOLIS IN 46226 USA Toll free: 800-374-8557 (in USA & Canada)
web: http://www.asmicro.com Fax: 317-895-5652
[business activities: analytical services in AFM, AFM probes, consulting,
training,
calibration and test specimens, calibration and measurement software,
used NanoScope equipment.] 2009-2010 is our 20th year in business!
=============================================
----- Original Message -----
From: John.Mardinly-at-asu.edu
Sent: Wednesday, April 04, 2012 2:58 PM

Bill, At that point it becomes a research project.

John Mardinly, ASU

On Apr 3, 2012, at 5:30 PM, "Bill & Sue Tivol" {wtivol-at-sbcglobal.net}
wrote:

}
} On Apr 3, 2012, at 12:49 PM, John.Mardinly-at-asu.edu wrote:
}
} } Marissa;
} } CMP means Chemical Mechanical Polish. That means that the silicon is
lightly etched while it is being polished. Any amorphization is so little
that it could not be detected by TEM. it is almost ready for growth of gate
oxide at this point. The epitaxial layer grown with silane is usually not
pure silane, but accompanied by some controlled impurities to dope the
surface of the wafer. You will never see any evidence in a TEM of the
interface between the epitaxial layer and the underlying wafer in
semiconductor quality silicon. The only thing you may see at the surface of
either CMP or epitaxial silicon would be a thin layer of oxide, called
native oxide. The thickness can depend on how long the silicon was exposed
to air and what other species may be adsorbed onto the surface. The native
oxide would generally be only a few angstroms thick.
} }
} } John Mardinly, ASU
} }
} } } Email: mlibbee-at-gmail.com Name: Marissa
} } }
} } } Organization: LBL- NCEM
} } }
} } } Title-Subject: [Filtered] Amorphous damage from CMP wafer preparation
} } }
} } } Message: During the Si wafer growth and preparation process, how much
damage is produced from the
} } } CMP process? Also, I've read that an epitaxial layer is grown on the
wafer by passing silane gas
} } } over the surface. Will someone comment on the way these processes
impact the structure of the Si at
} } } the surface of the wafer?
} } }
} } } Much obliged,
} } } Marissa
} } }
} Hi John and Marissa,
} Could grazing-incidence electron diffraction, used, for example, by
Larry Marks (sp?), detect and/or quantitate surface changes produced by CMP
and silane?
} Yours,
} Bill

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Hi everybody,

A colleague is going to buy an SEM; he is considering one from JEOL, Hitach= i or Tescan and asking for advice. The Tescan is a new company for me, so t= here is my question: how good is their service?

Thanks,

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

==============================Original Headers==============================
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Hi Nicolas Stephant,

ibss Group, Inc. offers the GV10x Asher in-situ plasma cleaner to reduce and
eliminate contamination in EM chambers and on specimens.

Please visit our web site, ibssGroup.com to learn about GV10x DS Asher
operation and contact us offline.

Vincent Carlino
ibss Group, Inc.
+1.415.566.5774
+1.415.566.9779 fax
vince.carlino-at-ibssgroup.com
www.ibssgroup.com

Commercial Disclaimer: ibss Group, Inc. is the manufacturer of the GV10x
in-situ plasma cleaner.

-----Original Message-----
X-from: Nicolas.Stephant-at-univ-nantes.fr
[mailto:Nicolas.Stephant-at-univ-nantes.fr]
Sent: Wednesday, April 04, 2012 1:19 AM
To: vcrvince-at-comcast.net

Dear listeners

I know only one system to clean samples inside a SEM to avoid contamination
dammage. Would you please inform if you know an alternate choice to EVACTRON
from XEI scientific ?
Thanks a lot.

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en produire
une image"

C.G Jung

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I am excited by a number of replies on my first question. Thanks a lot!

So, being still excited, I am posting my second for today question.
I am looking for a service laboratory for micro XRD work, to analyze small areas of specimen. I understand it is not proper question for Microscopy listserver, but at least both methods has "micro" in them.
Thanks,

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: DusevichV-at-umkc.edu [mailto:DusevichV-at-umkc.edu]
} Sent: Thursday, April 05, 2012 12:58 PM
} To: Dusevich, Vladimir
} Subject: [Microscopy] Tescan service
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
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} ----------------------------------------------------------------------------
}
} Hi everybody,
}
} A colleague is going to buy an SEM; he is considering one from JEOL, Hitach= i
} or Tescan and asking for advice. The Tescan is a new company for me, so t=
} here is my question: how good is their service?
}
} Thanks,
}
} Vladimir
}
} Vladimir M. Dusevich, Ph.D.
} Electron Microscope Lab Director
} 371 School of Dentistry
} 650 E. 25th Street
} Kansas City, MO 64108-2784
}
}
}
}
} ==============================Original
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All:
The Texas Society for Microscopy invites you to participate in
its 2012 meeting taking place on the campus of Texas Christian
University from April 12th - 14th.

Invited speakers include Dr. Ray H. Baughman of the Alan G.
MacDiarmid NanoTech Institute, The University of Texas at Dallas;
and Dr. Aydogan Ozcan of the Bio- and Nano-Photonics Laboratory,
UCLA. Workshops are scheduled for Thursday afternoon and include
"Sample Preparation Techniques for Materials Science - Choosing
the Right Method" by Robert Ranner, Product Manager, Leica
Applications Specialist.

The meeting location is the Brown-Lupton University Union while
the host hotel will be the SpringHill Suites by Marriott
University Fort Worth. Shuttle service will be available from the
hotel to campus . The abstract deadline is March 1. Additional
meeting information and forms may be found by visiting our
website (http://www.texasmicroscopy.org) or by emailing the
Secretary, Bob Droleskey, at bob.droleskey-at-ars.usda.gov.

We hope to see you there!

==============================Original Headers==============================
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Dear all,
We have an old Balzers 010 CPD for a long time. It just stopped to work after all those years of perfect work. The pressure chamber heating band has burnt off.
See: http://www2.biomed.cas.cz/~benada/CPD_failure.html

Please, does anybody knows if a replacement of the heating band still exists? I do not have any manual or scheme.

Thanking you in advance.

Happy Easter

Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

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7, 36 -- Subject: Balzers 010 CPD
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Hi Oldrich,

I am not familiar with Balzers 010 CPD and can't help you with finding
OEM spare part, but replacing flexible heater is a relatively
straight-forward task and any decent equipment technician should be able
to do it for you. In a nutshell you need to get following ifromation
about your heater:

1) Physical dimensions;
2. Voltage (can be measured on CPD unit);
3. Power rating or resistance - even with burned heater its resistance
can be measured and power rating estimated;
4. Temperature rating;

Once you have the dimensions, voltage, power, and temperature rating
then find a suitable replacement from what is currently available on the
market. Flexible heaters are sold by Omega, Watlow, Heatron, Minco and
quite a few other manufacturers, so I am sure that something suitable
for replacing your burned element can be found.

Good luck :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 4/6/2012 9:40 AM, benada-at-biomed.cas.cz wrote:
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} Dear all,
} We have an old Balzers 010 CPD for a long time. It just stopped to work after all those years of perfect work. The pressure chamber heating band has burnt off.
} See: http://www2.biomed.cas.cz/~benada/CPD_failure.html
}
} Please, does anybody knows if a replacement of the heating band still exists? I do not have any manual or scheme.
}
} Thanking you in advance.
}
} Happy Easter
}
} Oldrich
}
}

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Dear Luca,
I am just working on a SEM and found that some of the spikes (line distortions) might be generated by a mobile phone near by.
I would also check for mechanical vibrations first (do you have a turbo-molecular pump?) and then check all earth connections.

Best wishes,
Stefan

-------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D-97072 Wuerzburg Germany
Tel. +49-931-7848700
Fax +49-931-7848701
Mobile +49-175-7177051

Email: diller-at-stefan-diller.com
Websites: www.stefan-diller.com
www.elektronenmikroskopie.info
www. assisi.de
--------------------------------------------

Am 09.04.2012 um 14:53 schrieb microscopylistserver-noreply-at-microscopy.com:

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} Email: lfedele-at-vt.edu
} Name: Luca Fedele
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} Organization: Virginia Tech
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} Title-Subject: [Filtered] SEM - Image distortions
}
} Message: Dear All,
}
} We are experiences distortions/interferences (please check an example
} here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
} Camscan Series 2 through our Gatan Digiscan II interface.
}
} The images look ok on the old green phosphorous screens (but it might be
} because their resolution is limited of course) but come out distorted when
} captured.
}
} I have attached a BSE sample at high magnification (5000x) but the effect
} is also visible with SE images and at lower magnification (1000x or so).
} The effect remains regardless of software settings.
}
} We have several cables running around the SEM and suspect some sort of
} electrical noise (which could also be generated inside the computer
} connected via firewire to the digiscan) but so far we have not been able
} to find a cure.
}
} If anybody has ever seen an "interference" like this one and has ANY kind
} of suggestion we will be more than happy to give it a try :-)
}
} Thanks
}
} Luca
}
} -------------------------
} Luca Fedele PhD
} Department of Geosciences
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} Virginia Tech
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Oldrich,
I was given one of these with the same problem, plus the RTD for sensing
temperature was bad, if I recall. I haven't used it, but I did do a repair
that seems to do the trick.

The band heater is 25W and is supplied by a relay switched 250V. I bought a
very inexpensive 5A 5V switching power supply that had an indiscriminate
input requirement (something like 100-250VAC). I measured out enough
nichrome wire to give a 1 ohm resistance. I bought a can of high
temperature ceramic adhesive and put a layer on the outside of the chamber.
After it dried, I wrapped the nichrome wire around the chamber and put on
another layer of the ceramic adhesive and attached the wire to the 5V power
supply. Voila! A 25W heater operated by the switched 250V power source.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: benada-at-biomed.cas.cz [mailto:benada-at-biomed.cas.cz]
Sent: Friday, April 06, 2012 9:41 AM
To: kenconverse-at-qualityimages.biz

Dear all,
We have an old Balzers 010 CPD for a long time. It just stopped to work
after all those years of perfect work. The pressure chamber heating band has
burnt off.
See: http://www2.biomed.cas.cz/~benada/CPD_failure.html

Please, does anybody knows if a replacement of the heating band still
exists? I do not have any manual or scheme.

Thanking you in advance.

Happy Easter

Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

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Email: lfedele-at-vt.edu {mailto:lfedele-at-vt.edu}
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg
{http://www.lucafedele.eu/test1BSE5500.jpg} ) when capturing images
on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it
might be
because their resolution is limited of course) but come out
distorted when
captured.

I have attached a BSE sample at high magnification (5000x) but the
effect
is also visible with SE images and at lower magnification (1000x or
so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been
able
to find a cure.

If anybody has ever seen an "interference" like this one and has
ANY kind
of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu {mailto:lfedele-at-vt.edu}
www.lucafedele.eu {http://www.lucafedele.eu}
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--

Dr Nicholas Seaton

SEM Specialist
Characterization Facility

College of Science and Engineering

University of Minnesota

Shepherd Labs

100 Union Street SE

Minneapolis

MN, 55455

email: seato008-at-umn.edu {mailto:seato008-at-umn.edu}

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You say that is a BSE image. The brightness variation is a bit much for a BSE image. Since I see you are with the department of geosciences, I suspect charging is present. It may be contributing to the appearance as much as any electrical interference.

I would suggest two things.

First, I would recommend cutting back on current if it is higher than normal. If you are truly doing everything else (coating, grounding the sample. voltage, etc) the same, then high current may be leading to charging. I presume you have EDS on this system. What kind of total count rate do you get under these conditions?

Second, I would just put this sample aside and examine something conductive like a block of metal or a sample holder. If it shows distortion, then you know you have some other problem to track down.

Warren Straszheim
________________________________________
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Email: lfedele-at-vt.edu
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it might be
because their resolution is limited of course) but come out distorted when
captured.

I have attached a BSE sample at high magnification (5000x) but the effect
is also visible with SE images and at lower magnification (1000x or so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been able
to find a cure.

If anybody has ever seen an "interference" like this one and has ANY kind
of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
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...
Just another idea:
Does your SEM have some filters in the signal-path which change bandwith with the configuration of line / frame settings? It might
also be a wrong filter bandwith contributing to this kind of signal if your image aquisition system does not have his own scan
generator...

Best wishes,
Stefan

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

Am 09.04.12 14:53, schrieb microscopylistserver-noreply-at-microscopy.com:
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} Email: lfedele-at-vt.edu
} Name: Luca Fedele
}
} Organization: Virginia Tech
}
} Title-Subject: [Filtered] SEM - Image distortions
}
} Message: Dear All,
}
} We are experiences distortions/interferences (please check an example
} here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
} Camscan Series 2 through our Gatan Digiscan II interface.
}
} The images look ok on the old green phosphorous screens (but it might be
} because their resolution is limited of course) but come out distorted when
} captured.
}
} I have attached a BSE sample at high magnification (5000x) but the effect
} is also visible with SE images and at lower magnification (1000x or so).
} The effect remains regardless of software settings.
}
} We have several cables running around the SEM and suspect some sort of
} electrical noise (which could also be generated inside the computer
} connected via firewire to the digiscan) but so far we have not been able
} to find a cure.
}
} If anybody has ever seen an "interference" like this one and has ANY kind
} of suggestion we will be more than happy to give it a try :-)
}
} Thanks
}
} Luca
}
} -------------------------
} Luca Fedele PhD
} Department of Geosciences
} 4044 Derring Hall
} Virginia Tech
} Blacksburg, VA, 24061
} lfedele-at-vt.edu
} www.lucafedele.eu
} ----------------------
}
} Login Host: 204.111.131.89
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An EM facility is being designed locally, and one consideration is fire protection. A bit of research reveals that some IT and telecommunications facilities have opted either for Dupont's "FM-200" heat absorption chemistry, or gaseous "Intergen" which is a smothering agent. Both are applicable for use in normally occupied areas.

A search of the Microscopy archives revealed little in the way of recent discussions ... does anyone have first hand experience with either of these methods? ... or would anyone care to suggest something different???

Cheerios from "The Rock" ...
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Genuinely, Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada

cogito ergo ZzoooomM

This electronic communication is governed by the terms and conditions at
http://www.mun.ca/cc/policies/electronic_communications_disclaimer_2011.php

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The Company:
Our client is a leading developer and producer of innovative high-tech
precision optics systems for the analysis of microstructures. As one of
the market leaders in each of the fields of Microscopy, Confocal Laser
Scanning Microscopy, Imaging Systems, Specimen Preparation and Medical
Equipment. Comprising nine manufacturing facilities in seven countries,
sales and service companies in 20 countries and an international network
of dealers, the company is represented in over 100 countries.

The Opportunity:
The company currently has an opening for a Microscopy Sales Representative
to be based in Dallas TX. All applicants must not be adverse to travel,
as this is a position that may require you to travel when necessary.

Base: Commensurate with experience

Other: Full benefits - 401k program/matching - Car allowance - Company
cell phone - Laptop computer - Home office set-up - Expenses

Primary Responsibilities:
As a key member of the sales team, the purpose of this position is to
promote sales of the company's product line in the assigned region. The
primary function is to execute domestic sales plans, recommend strategies
to improve the sale of products in the territory and aid in the
implementation of these strategies in line with Marketing, Corporate
growth, profitability and mission objectives.

Additional Responsibilities:
- Schedule product seminars or training sessions in concert with company
Product Management in the assigned region
- Participate in all activities that will enhance the awareness,
acceptance and sale of products
- Conduct analysis and performance evaluation of all sales by use of the
company's Customer Resource Management tool
- Work with Sales Manager to develop strategies designed to increase sales
of products; Specifically, the Microscopy Sales Representative will
recommend product, pricing, advertising and sales promotion strategies
- Coordinates the generation of new product ideas or modifications,
performs analysis and preliminary screening of these ideas and recommends
the appropriate ones to the Area Sales Manager for acceptance or rejection

Education and Experience Required:
BA/BS or MS degree in physics or chemistry or a related scientific field
is highly preferred. Experience working with microscopy products is
required and a minimum of 2 years related microscopy experience in a
laboratory is essential. In-depth knowledge of the microscopy market is
also highly desirable. Sales experience in selling products of a
technical nature is desirable but not mandatory. The individual must be
highly numerate as pricing, quoting and reviewing commercial terms will be
a daily routine. Strong computer skills and experience with CRM systems
(Maximizer, ACT, Goldmine, Siebel, etc). Strong technical,
electrochemical, optical and mechanical aptitude.

If you meet and/or exceed the experience criteria, please submit your
resume by emailing Christy Edwards (Sr. e-Recruitment Consultant with
Personify) at ce-at-personifysearch.com. We wish everyone the best of luck.
Unfortunately only qualified candidates will be considered.

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X-from: Nick Seaton {seato008-at-umn.edu}
To: microscopylistserver-noreply-at-microscopy.com

Luca,

It really looks like mechanical vibration to me. I have seen this kind of noise in images when the
scope is being subjected to mechanical vibrations, but also when the sample is not well clamped down
inside the chamber the beam can cause sample movement too resulting in vibrations along the edges of
objects in the image. Although this usually requires much higher mags to see.

Nick Seaton

On Mon, Apr 9, 2012 at 8:00 AM, {microscopylistserver-noreply-at-microscopy.com
{mailto:microscopylistserver-noreply-at-microscopy.com} } wrote:

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Email: lfedele-at-vt.edu {mailto:lfedele-at-vt.edu}
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg {http://www.lucafedele.eu/test1BSE5500.jpg} ) when
capturing images on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it might be
because their resolution is limited of course) but come out distorted when
captured.

I have attached a BSE sample at high magnification (5000x) but the effect
is also visible with SE images and at lower magnification (1000x or so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been able
to find a cure.

If anybody has ever seen an "interference" like this one and has ANY kind
of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu {mailto:lfedele-at-vt.edu}
www.lucafedele.eu {http://www.lucafedele.eu}
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--

Dr Nicholas Seaton

SEM Specialist
Characterization Facility

College of Science and Engineering

University of Minnesota

Shepherd Labs

100 Union Street SE

Minneapolis

MN, 55455

email: seato008-at-umn.edu {mailto:seato008-at-umn.edu}

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Email: lfedele-at-vt.edu
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it might be
because their resolution is limited of course) but come out distorted
when captured.

I have attached a BSE sample at high magnification (5000x) but the
effect is also visible with SE images and at lower magnification (1000x
or so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been able
to find a cure.

If anybody has ever seen an "interference" like this one and has ANY
kind of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
----------------------

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Email: lfedele-at-vt.edu
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it might be
because their resolution is limited of course) but come out distorted
when captured.

I have attached a BSE sample at high magnification (5000x) but the
effect is also visible with SE images and at lower magnification (1000x
or so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been able
to find a cure.

If anybody has ever seen an "interference" like this one and has ANY
kind of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
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Hi Luca

To judge exactly what is happening you need to help us by providing more
information; one picture does not tell the full story!

Use the standard fault finding procedure - "Is the "fault" the same at short
and long working distances?"

If so it is almost certainly a vibration, if not then it is almost certainly
a magnetic field. Vibration should not change no matter what the working
distance or accelerating voltage. Magnetic fields are reduced at higher
accelerating voltage or at very short working distances the specimen being
protected by the higher lens fields.

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
www.emcourses.com

} Email: lfedele-at-vt.edu
} Name: Luca Fedele
}
} Organization: Virginia Tech
}
} Title-Subject: [Filtered] SEM - Image distortions
}
} Message: Dear All,
}
} We are experiences distortions/interferences (please check an example
} here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
} Camscan Series 2 through our Gatan Digiscan II interface.
}
} The images look ok on the old green phosphorous screens (but it might be
} because their resolution is limited of course) but come out distorted when
} captured.
}
} I have attached a BSE sample at high magnification (5000x) but the effect
} is also visible with SE images and at lower magnification (1000x or so).
} The effect remains regardless of software settings.
}
} We have several cables running around the SEM and suspect some sort of
} electrical noise (which could also be generated inside the computer
} connected via firewire to the digiscan) but so far we have not been able
} to find a cure.
}
} If anybody has ever seen an "interference" like this one and has ANY kind
} of suggestion we will be more than happy to give it a try :-)
}
} Thanks
}
} Luca
}
} -------------------------
} Luca Fedele PhD
} Department of Geosciences
} 4044 Derring Hall
} Virginia Tech
} Blacksburg, VA, 24061
} lfedele-at-vt.edu
} www.lucafedele.eu
} ----------------------
}
} Login Host: 204.111.131.89
} ---------------------------------------------------------------------------
}
} ===========================================
} Do not reply to this message it is from
} the Microscopy Listserver NO-REPLY forwarding
} system. You should send a new message to
}
} Microscopy-at-Microscopy.com
}
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12, 26 -- From protrain-at-emcourses.com Tue Apr 10 07:58:50 2012
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Hi Luca,

Checking the effect of accelerating voltage is a good idea to ensure
magnetic field or not. One way to check vibration is to design an
assembly to fix a sample to the lens (as a backscattered detector can
be) and NOT to the stage. Like that, if there is vibration, sample and
beam are synchronized and you should not see effect on the image.
I would be very interesting to check carefully the green screen to know
if the aberration is same or not (by increasing magnification ?).
Regards

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

-------- Message original --------
Sujet: [Microscopy] Re: viaWWW:SEM Image distortions

Hi Luca

To judge exactly what is happening you need to help us by providing more
information; one picture does not tell the full story!

Use the standard fault finding procedure - "Is the "fault" the same at short
and long working distances?"

If so it is almost certainly a vibration, if not then it is almost certainly
a magnetic field. Vibration should not change no matter what the working
distance or accelerating voltage. Magnetic fields are reduced at higher
accelerating voltage or at very short working distances the specimen being
protected by the higher lens fields.

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
www.emcourses.com

} Email: lfedele-at-vt.edu
} Name: Luca Fedele
}
} Organization: Virginia Tech
}
} Title-Subject: [Filtered] SEM - Image distortions
}
} Message: Dear All,
}
} We are experiences distortions/interferences (please check an example
} here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
} Camscan Series 2 through our Gatan Digiscan II interface.
}
} The images look ok on the old green phosphorous screens (but it might be
} because their resolution is limited of course) but come out distorted when
} captured.
}
} I have attached a BSE sample at high magnification (5000x) but the effect
} is also visible with SE images and at lower magnification (1000x or so).
} The effect remains regardless of software settings.
}
} We have several cables running around the SEM and suspect some sort of
} electrical noise (which could also be generated inside the computer
} connected via firewire to the digiscan) but so far we have not been able
} to find a cure.
}
} If anybody has ever seen an "interference" like this one and has ANY kind
} of suggestion we will be more than happy to give it a try :-)
}
} Thanks
}
} Luca
}
} -------------------------
} Luca Fedele PhD
} Department of Geosciences
} 4044 Derring Hall
} Virginia Tech
} Blacksburg, VA, 24061
} lfedele-at-vt.edu
} www.lucafedele.eu
} ----------------------
}
} Login Host: 204.111.131.89
} ---------------------------------------------------------------------------
}
} ===========================================
} Do not reply to this message it is from
} the Microscopy Listserver NO-REPLY forwarding
} system. You should send a new message to
}
} Microscopy-at-Microscopy.com
}
} ============================================
}
}
}
} ==============================Original
} Headers==============================
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} 07:47:33 2012
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26, 23 -- From Nicolas.Stephant-at-univ-nantes.fr Tue Apr 10 08:33:58 2012
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after many years of testing variations, I came up to the following
schedule which works fine (at least most of the time):

place section (usually with fine forceps) on a drop of 20% acetone in
distilled water on SuperfrostPlus glass slides (yes, the really
expensive ones from Menzel Glass) and let dry at 60� (Celsius), after
complete drying (no blowing or fanning air to speed up drying) transfer
to hot plate with 80�, wait two minutes and stain at 80� with
Richardson's Blue.
I use precision hot plates that do not oscillate in temperature. The
magnetic heat-stirrers are in my opinion useless due to strongly
oscillating temperature leading to gas bubbles under the section.
I usually cut Araldite at 1,5 or 2,5 �m with a glass knife. With respect
to "wrinkles", Diamond histo-knifes seem not to be the improvement.
Trimming down the sectional area as far as possible is of course
advantageous. If from time to time everything fails, I employ degassed
water (or freshly from the destille) with acetone.
In my opinion, the"knack" is to use 20% acetone and dry at no more than
60� (Celsius).

good luck,

Peter

PS: for those who can read German: there is a fine book by Peter B�ck
called "Der Semid�nnschnitt" ("the semi-thin section") published in the
mid-eighties.

--
====================================================

Dr. Peter Heimann Raum: W7-107 / Tel.: +49-(0)521-106-5628
Universitaet Bielefeld Fax: +49-(0)521-106-5654
"Zellbiologie / Cell Biology" W7-107
33501 Bielefeld, Germanywww.uni-bielefeld.de/biologie/cellbio

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Hi Luca,

You see the fuzzy edges along the grain boundary is likely caused by the
mechanical vibration interferences. You would see this effect with any
other detectors if the vibration source remains. The problem will be
resolved once the vibration source is spotted and removed.

Make sure no external object (e.g. table) is in direct contact with SEM main
console. Cables need to loosely lay on the ground. It is also a good idea
to put a rubber mat under the rotary pump.

Regards
---------------------------------
Xiang Yang
Regional Analytical Centre
Saint Mary's University
Halifax, NS Canada B3H 3C3
Phone: 902-420-5709
http://www.smu.ca/institutes/emc/welcome.html
http://www.smu.ca/institutes/rgc/welcome.html

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Email: lfedele-at-vt.edu
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it might be
because their resolution is limited of course) but come out distorted when
captured.

I have attached a BSE sample at high magnification (5000x) but the effect
is also visible with SE images and at lower magnification (1000x or so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been able
to find a cure.

If anybody has ever seen an "interference" like this one and has ANY kind
of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
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Hi Luca,

The interference that you have in your image looks pretty periodic. Have you
tried to measure the periodicity? The periodicity is often a good point to
start. If it is 50 or 60 Hz, it is most likely an electrical problem, which
could be due to ground loops. In that case I would suggest to connect the
entire acquisition system to the same power source and ground as the
microscope. I have also seen this when the start of the line acquisition is
"at random" with respect to the phase of the power signal. You may want to
synchronize the start of the line acquisition with the phase of the power
signal.

If the periodicity is not 50 or 60 Hz, I would check out the sources for
noise already mentioned here. You may also want to check out any electrical
equipment that is nearby. A colleague of mine found out after some sleuthing
that the spurious signal he had in his data came from an elevator quite a
distance away. They ended up blocking the elevator when they had to make
sensitive measurements...

Mike Bode, Ph.D.
ResAlta Research Technologies Corp.

2102 Beech Ct
Golden, CO 80401
USA

p: +1-303-234-9270
f: +1-303-202-6350
Mike.Bode-at-ResAltaTech.com
www.ResAltaTech.com

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Email: lfedele-at-vt.edu
Name: Luca Fedele

Organization: Virginia Tech

Title-Subject: [Filtered] SEM - Image distortions

Message: Dear All,

We are experiences distortions/interferences (please check an example
here: www.lucafedele.eu/test1BSE5500.jpg) when capturing images on our
Camscan Series 2 through our Gatan Digiscan II interface.

The images look ok on the old green phosphorous screens (but it might be
because their resolution is limited of course) but come out distorted when
captured.

I have attached a BSE sample at high magnification (5000x) but the effect is
also visible with SE images and at lower magnification (1000x or so).
The effect remains regardless of software settings.

We have several cables running around the SEM and suspect some sort of
electrical noise (which could also be generated inside the computer
connected via firewire to the digiscan) but so far we have not been able to
find a cure.

If anybody has ever seen an "interference" like this one and has ANY kind
of suggestion we will be more than happy to give it a try :-)

Thanks

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
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Dear list,

I have a strange behavior during a LM fixation and I would appreciate your feedback.
I prepared my fixative as follows:
- suspended 2 grams of a 6-years old paraformaldehyde powder in mQ water
-Heated at 60�C for 10 min with magnetic stirring.
- Added 3 drops of 1N NaOH to depolymerize
- let cool down, then double the volume with 100mM Hepes pH 7.2
Final solution is 4% PFA in 50mM Hepes pH 7.2

I used the solution 2 hours after preparation on cells grown on coverslips. I washed them just once with PBS before adding fixative.
Under the LM I can see impressive blebbing of the cell membrane and that's my concern.
It seems that my solution is hypoosmotic, is it the case?
In this hypothesis I could just add some NaCl to make it isotonic. At the bottom of my memory I seem to remember that isoosmoticity sould be approx. 260mosm.
The problem is that I don't know how to take account of the�presence of formaldehyde for the calculation of the osmolarity.
At the �beginnning of the fixation I suppose that only a few molecules cross the membrane but as the fixation proceeds more and more molecules cross the membrane. 4% is quite high so if�it counts I guess that the solution is not hypoosmotic.

Stephane

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Hello,
I would like to thank everybody who send me an advice, hint or suggestion how to solve the problem with our CPD. I've even got a manual with scheme for this device from local resources, although in German.

My thanks and best regards

Oldrich
--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic
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} Dear all,
} We have an old Balzers 010 CPD for a long time. It just stopped to work
} after all those years of perfect work. The pressure chamber heating band has
} burnt off.
} See: http://www2.biomed.cas.cz/~benada/CPD_failure.html
}
} Please, does anybody knows if a replacement of the heating band still
} exists? I do not have any manual or scheme.
}
} Thanking you in advance.
}
} Happy Easter
}
} Oldrich
}
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3, 38 -- From benada-at-biomed.cas.cz Wed Apr 11 05:58:34 2012
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Blebbing in formaldehyde is a well-known phenomenon. I think Elizabeth Hay's group may have been the first to describe. Adding a small amount of glutaraldehyde can prevent. Formaldehyde penetrates quickly but reacts more slowly than glutaraldehyde. Osmolarity is only one factor. You are permeabilizing the membrane and allowing calcium and other electrolytes to cross. The only real solution is quick-freezing.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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To: Phillips, Thomas E.

Dear list,

I have a strange behavior during a LM fixation and I would appreciate your feedback.
I prepared my fixative as follows:
- suspended 2 grams of a 6-years old paraformaldehyde powder in mQ water -Heated at 60�C for 10 min with magnetic stirring.
- Added 3 drops of 1N NaOH to depolymerize
- let cool down, then double the volume with 100mM Hepes pH 7.2 Final solution is 4% PFA in 50mM Hepes pH 7.2

I used the solution 2 hours after preparation on cells grown on coverslips. I washed them just once with PBS before adding fixative.
Under the LM I can see impressive blebbing of the cell membrane and that's my concern.
It seems that my solution is hypoosmotic, is it the case?
In this hypothesis I could just add some NaCl to make it isotonic. At the bottom of my memory I seem to remember that isoosmoticity sould be approx. 260mosm.
The problem is that I don't know how to take account of the�presence of formaldehyde for the calculation of the osmolarity.
At the �beginnning of the fixation I suppose that only a few molecules cross the membrane but as the fixation proceeds more and more molecules cross the membrane. 4% is quite high so if�it counts I guess that the solution is not hypoosmotic.

Stephane

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5, 36 -- Date: Wed, 11 Apr 2012 03:40:23 -0700 (PDT) 5, 36 -- From: Stephane Nizet {nizets2-at-yahoo.com} 5, 36 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com} 5, 36 -- Subject: Fixation for LM 5, 36 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com} 5, 36 -- MIME-Version: 1.0 5, 36 -- Content-Type: text/plain; charset=iso-8859-1 5, 36 -- Content-Transfer-Encoding: 8bit 5, 36 -- X-MIME-Autoconverted: from quoted-printable to 8bit by ns.microscopy.com id q3BAeOFL007009 ==============================End of - Headers==============================

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14, 31 -- From PhillipsT-at-missouri.edu Wed Apr 11 07:55:10 2012
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Dear Stephane,

it is already known (or at least has been described) for long that the fixative itself - despite perhaps hyperosmotic - does not contribute much to tonicity of the fixation process, instead, the tonicity/osmolarity/osmolality of the fixative vehicle = buffer solution is decisive
(e.g. Willen H, Willen R & Stendahl U: Invasive squamous cell carcinoma of the uterine cervix. VII. Influence of vehicle osmolarity on biopsy quality. Acta Radiologica 22 No 4,257-261,1983).

Other references - surprisingly -(e.g. Margo CE, Lee A.: Fixation of whole eyes: the role of fixative osmolarity in the production of tissue artifact. Graefes Arch Clin Exp Ophthalmol. 1995 Jun;233(6):366-70.) tell us the contrary.

(NB: these references are not the only ones for sure, but I had them at hand right now)

HEPES (a zwitterionic buffer) used in /as the medium at a concentration of 0.01 M (= 100 mM) showed adequate buffering capacity and was not cytotoxic to monolayer cultures (cf. BRIMBLE TW: The developing of the zwitterionic buffer; KONTAKTE (Merck?) 1981(No1) p. 37-43) other references state: "no apparent cytotoxicity when HEPES was used up to 28 mM

So, if I remember correctly the original Karnovsky fixative exerts hyperosmolarity in its component formaldehyde (around 2000 mosmol)
(cf. Karnovsky (M.J. Karnovsky. 1965. A formaldehyde-glutaraldehyde fixative of high osmolality for use in electron microscopy. J. Cell Biol. 27:137A): sodium cacodylate-buffered (80mM), calcium chloride-enhanced (5 mM) mixture of 5% glutaraldehyde and 4% formaldehyde (prepared from paraformaldehyde powder).
The original recipe produces a hypertonic medium (2010 mOsM) that was so hypertonic that most investigators made it up at {half-strength} .

Osmolarity of a 4% hydrous Formaldehyde solution is (calculated chemically) 1,333 mOsmol (IMHO this is quite a high value for cells on a coverslip).

Isoosmoticity (isotonicity) of blood is regarded/reported to be +/- 300 mosmol (for PO4 buffer = approx. 0.13M), but I have measured commercially available {physiological saline} as low as 260 mosmol).
But there are several kinds of tissues (e.g. liver cell isotonicity = 0.34M, kidney [cells] 0.23M etc.) which obviousely by chemical fixation would be preserved better using osmolarities of fixative solutions differing from the {blood isotonic} osmolarity (cf. Millonig G(C): Laboratory Manual of Biological Electron microscopy, pp1-64, 1976 Mario Saviolo-Editore Vercelli C.P. 182, Italy).

Also it might be of interest which buffer system you use to get the correct ionic strength or ionic pressure which does not interfere with the preservation of cells as an {aequivalent image} as is called the state of the art (chemical) fixation process. So Millonig and other references also tell that using usual buffer solution with formaldehyde you'll end up with 160 mosmol, if you use the Millonigs Na-NA-PO4-buffer, youl end up with 290 msomol which is nearly at the isotonic value for blood.

You haven't said which kind of cells you are treating on the coverslips. I guess these cells are monolayers and much more vulnerable to fixation artifacts than bioptic tissue. SO also fixation time and temperature perhaps counts. So it might advisable to compare your recipe with the literature for similar cell layer preps.

Pure 25% GA I've measured to have approx. 1700 mosmol, 12.5% 853, 1% around 67-69 and 0.5% 28-29 mosmol (with the latter two measurements you can see that diluting 1% GA one:one will not result in the half percentage! which is perhaps due to [ionic?] dissociation phenomena).
Dissociation phenomena also contribute to the fact that diluting 0.2 M Phosphate buffer 1:1 with A. dest. will not result in an osmol-value which is half that of 0.2 M (it is generally a little bit higher)

LM-Fixatives (you can find those simply by googeling):
e.g. "McDowell's and Trump's 4F:1G": This fixative contains 4% formaldehyde and 1% glutaraldehyde in a phosphate buffer, with an osmolarity of 176 mOsM (unfortunately, the values for the fixatives and buffers are not given separately)
Contrary to that, a "Modified 4F:1G Recipe" (same source!) states: 4% formaldehyde/1% glutaraldehyde fixative in 0.1 M sodium phosphate buffer utilizing paraformaldehyde powder; The final pH should be approximately 7.3 (check it), and the osmolarity approximately 1583 mOsM.

Conc. the blebbing of your cells: does this appear also when flooding the coverslips with PBS? (which might also not be isotonic to the cells' fluid.

There are a lot of recipes out there (many of them in my files)
hope you'll find the right one

best wishes and good luck,

Wolfgang MUSS PhD
SALZBURG, Austria

} -----Urspr�ngliche Nachricht-----
} Von: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Gesendet: Mittwoch, 11. April 2012 12:44
} An: Mu� Wolfgang
} Betreff: [Microscopy] Fixation for LM
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} Dear list,
}
} I have a strange behavior during a LM fixation and I would appreciate
} your feedback.
}
} I prepared my fixative as follows:
} - suspended 2 grams of a 6-years old paraformaldehyde powder in mQ water
} -Heated at 60�C for 10 min with magnetic stirring.
} - Added 3 drops of 1N NaOH to depolymerize
} - let cool down, then double the volume with 100mM Hepes pH 7.2
} Final solution is 4% PFA in 50mM Hepes pH 7.2
}
} I used the solution 2 hours after preparation on cells grown on
} coverslips. I washed them just once with PBS before adding fixative.
} Under the LM I can see impressive blebbing of the cell membrane and
} that's my concern.
} It seems that my solution is hypoosmotic, is it the case?
} In this hypothesis I could just add some NaCl to make it isotonic. At
} the bottom of my memory I seem to remember that isoosmoticity should be
} approx. 260mosm.
} The problem is that I don't know how to take account of the�presence of
} formaldehyde for the calculation of the osmolarity.
} At the �beginnning of the fixation I suppose that only a few molecules
} cross the membrane but as the fixation proceeds more and more molecules
} cross the membrane. 4% is quite high so if�it counts I guess that the
} solution is not hypoosmotic.
}
} Stephane
}
}
} ==============================Original
} Headers==============================
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I use 10% of the Formaldehyde molarity and 30% of the Glut molarity and add that to the buffer. This seems to work well for total.
David

On Apr 11, 2012, at 3:42 AM, {nizets2-at-yahoo.com} {nizets2-at-yahoo.com} wrote:

}
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} Dear list,
}
} I have a strange behavior during a LM fixation and I would appreciate your feedback.
} I prepared my fixative as follows:
} - suspended 2 grams of a 6-years old paraformaldehyde powder in mQ water
} -Heated at 60�C for 10 min with magnetic stirring.
} - Added 3 drops of 1N NaOH to depolymerize
} - let cool down, then double the volume with 100mM Hepes pH 7.2
} Final solution is 4% PFA in 50mM Hepes pH 7.2
}
} I used the solution 2 hours after preparation on cells grown on coverslips. I washed them just once with PBS before adding fixative.
} Under the LM I can see impressive blebbing of the cell membrane and that's my concern.
} It seems that my solution is hypoosmotic, is it the case?
} In this hypothesis I could just add some NaCl to make it isotonic. At the bottom of my memory I seem to remember that isoosmoticity sould be approx. 260mosm.
} The problem is that I don't know how to take account of the presence of formaldehyde for the calculation of the osmolarity.
} At the beginnning of the fixation I suppose that only a few molecules cross the membrane but as the fixation proceeds more and more molecules cross the membrane. 4% is quite high so if it counts I guess that the solution is not hypoosmotic.
}
} Stephane
}
}
} ==============================Original Headers==============================
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Listers:

I just received the following request from Lee Gould ("Lee Gould"
{lgould-at-nmu.edu} ) and thought it worthwhile forwarding it to you-all:

: " I found a website with RCA microscope information
on it and your name and contact information was provided. I am
looking for some help with RCA EMU-3G and EMU-4. Specifically
Northern Michigan University has one each of these and would like to
get rid of them. I see they could contain Radiation? Any other
hazardous materials? Any one interested in these microscopes? Not
sure you can help, but any information you can provide would be much
appreciated, or any contact information to anyone who can help?
Thank you for your valuable time!"

The RCA EMU-3 was built in the mid 1950s, and the EMU-4 in the mid
1960s, and so both instruments might well be considered "antiques".
I doubt that anyone would be interested in trying to get either back
into operation; however, maybe they would be valuable as a
collector's item or for a museum display.

--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

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Thanks for the replies.
Most answers point out that swelling of the cells is a known effect of formaldehyde fixation and that increasing the osmolarity of fixative could be favorable.
Some say that the concentration of formaldehyde should not be taken into account for any osmotic effect, one person uses only 10% of the formaldehyde concentration for the calculation.
�
Now a new but related question: I have found a 6-years old bottle of 37% formalin which was already open (but maintained closed) and stored at RT.
Do you think it is still good for basic IF work? I just want to stain one cell type in a mix culture so it is no high precision work here.
�
Stephane
�
�
�
�
} � ----------------------------------------------------------------------------
} � The Microscopy ListServer -- CoSponsor:� The Microscopy Society of America
} � To� Subscribe/Unsubscribe --
} � On-Line Help
} � ----------------------------------------------------------------------------
} �
} � Dear list,
} �
} � I have a strange behavior during a LM fixation and I would appreciate
} your feedback.
} � I prepared my fixative as follows:
} � - suspended 2 grams of a 6-years old paraformaldehyde powder in mQ water
} � -Heated at 60�C for 10 min with magnetic stirring.
} � - Added 3 drops of 1N NaOH to depolymerize
} � - let cool down, then double the volume with 100mM Hepes pH 7.2
} � Final solution is 4% PFA in 50mM Hepes pH 7.2
} �
} � I used the solution 2 hours after preparation on cells grown on
} coverslips. I washed them just once with PBS before adding fixative.
} � Under the LM I can see impressive blebbing of the cell membrane and
} that's my concern.
} � It seems that my solution is hypoosmotic, is it the case?
} � In this hypothesis I could just add some NaCl to make it isotonic. At
} the bottom of my memory I seem to remember that isoosmoticity sould be
} approx. 260mosm.
} � The problem is that I don't know how to take account of the�presence
} of formaldehyde for the calculation of the osmolarity.
} � At the �beginnning of the fixation I suppose that only a few
} molecules cross the membrane but as the fixation proceeds more and
} more molecules cross the membrane. 4% is quite high so if�it counts I
} guess that the solution is not hypoosmotic.
} �
} � Stephane
} �
} �
} � ==============================Original Headers==============================
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2, 43 -- Date: Thu, 12 Apr 2012 01:48:17 -0700 (PDT)
2, 43 -- From: Stephane Nizet {nizets2-at-yahoo.com}
2, 43 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com}
2, 43 -- Subject: Re: [Microscopy] Fixation for LM- Thanks and another question
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Dear Stephane,

thanks for at least resuming about other answers you got off-list concerning your first question.

I think it is worth to know about the "differences" between "osmolarity" and "tonicity" (which is/has been discussed not only once, also on this MSA listserver).

Concerning your "new" question:

You don't say anything about brand or storage conditions of the bottle of 37% Formalin
(guessing - for Austria - it might be the MERCK Formalin 37% [Histological grade or p. A.[pro analysi]-grade]
103999 Formaldehyde solution min. 37% free from acid
stabilized with about 10% methanol and calcium carbonate for histology
[= http://www.chemdat.merck.de/singapore/formaldehyde-solution-min-37-percent-free-from-acid/MDA_CHEM-103999/p_b92b.s1Ld8QAAAEWY.EfVhTl = ]

The stabilizer Methanol as well as the added calcium carbonate function in prevention of polymerization and as buffer medium for generated (formic) acid.

Usually (former times) the product had imprinted an expiration date on the label, which isn't always followed today,
e.g. a fresh bottle of that product was purchased in 2010-10 and show(s)/showed an expiration date 2012-05.

Assuming you can see that expiration date - under certification rules - one can / is allowed to use the content until 2012-05 without any consequences. If you / your lab aren't/isn't subject to such regulations I would consider the use of that charge/lot of fluid (if storage was in agreement with the suppliers data) until End of 2012 is o.k. After that date one could use the solution for fixation e.g. in Routine Histology/fixation of corpses etc.

Last but not least: you don't know about the real concentration as well as about the grade of polymerization in your old solution. Knowing that {filtering} does not really withdraw polymers of FA, one should be careful in pipetting or spilling the solution out of the bottle...latter perhaps may churn up the calcium carbonate powder which perhaps interferes with your microscopical preparatory intentions.

Best regards,
Wolfgang MUSS
EM-Lab, Pathology
SALK-LKH (Gen. Hosp.)
SALZBURG-Austria

Disclaimer: no financial interest in MERCK products - only user of such.

} -----Urspr�ngliche Nachricht-----
} Von: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Gesendet: Donnerstag, 12. April 2012 10:52
} An: Mu� Wolfgang
} Betreff: [Microscopy] Re: Fixation for LM- Thanks and another question
}
} ---------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of
} America
} To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Thanks for the replies.
} Most answers point out that swelling of the cells is a known effect of
} formaldehyde fixation and that increasing the osmolarity of fixative
} could be favorable.
} Some say that the concentration of formaldehyde should not be taken
} into account for any osmotic effect, one person uses only 10% of the
} formaldehyde concentration for the calculation.
}
} Now a new but related question: I have found a 6-years old bottle of
} 37% formalin which was already open (but maintained closed) and stored
} at RT.
} Do you think it is still good for basic IF work? I just want to stain
} one cell type in a mix culture so it is no high precision work here.
}
} Stephane
}
}
}
} } � --------------------------------------------------------------------
} --------
} } � The Microscopy ListServer -- CoSponsor:� The Microscopy Society of
} America
} } � To� Subscribe/Unsubscribe --
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} --------
} }
} } � Dear list,
} }
} } � I have a strange behavior during a LM fixation and I would
} appreciate
} } your feedback.
} } � I prepared my fixative as follows:
} } � - suspended 2 grams of a 6-years old paraformaldehyde powder in mQ
} water
} } � -Heated at 60�C for 10 min with magnetic stirring.
} } � - Added 3 drops of 1N NaOH to depolymerize
} } � - let cool down, then double the volume with 100mM Hepes pH 7.2
} } � Final solution is 4% PFA in 50mM Hepes pH 7.2
} }
} } � I used the solution 2 hours after preparation on cells grown on
} } coverslips. I washed them just once with PBS before adding fixative.
} } � Under the LM I can see impressive blebbing of the cell membrane and
} } that's my concern.
} } � It seems that my solution is hypoosmotic, is it the case?
} } � In this hypothesis I could just add some NaCl to make it isotonic.
} At
} } the bottom of my memory I seem to remember that isoosmoticity sould
} be
} } approx. 260mosm.
} } � The problem is that I don't know how to take account of the�presence
} } of formaldehyde for the calculation of the osmolarity.
} } � At the �beginnning of the fixation I suppose that only a few
} } molecules cross the membrane but as the fixation proceeds more and
} } more molecules cross the membrane. 4% is quite high so if�it counts I
} } guess that the solution is not hypoosmotic.
} }
} } � Stephane
} }
} }
} } � ==============================Original
} Headers==============================
} } � 5, 36 -- From nizets2-at-yahoo.com Wed Apr 11 05:40:24 2012
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Contents Retrieved from Microscopy Listserver Archives
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Talk about a trip down memory lane. When I joined Goodyear Tire they had a RCA EUM-4. It was just celebrating it's 25 service anniversary. We photographed a GY service pin in B&W, tinted it yellow and pasted it on the column. We used that scope for several more years before it failed and could not be resurrected no matter what the resourceful RCA technicians could do. We replaced it with Phillips 300, a great TEM, but it never had the romance of seeing ultra small particles for the first time like the RCA had.

I always wanted to put that TEM in the GY Rubber Museum complete with nice photomicrographs of carbon black, clay and other fillers used in tires, but who listens to worker bees. Even that museum is gone now.

Before that GY had a RCA-3, I've only seen photos or line drawings of it. The original microscopist at GY, Dr. Wilsferd (I sure I'm butchering his name). He was retired by the time I got there, but when he was in town he would stop in. He did some interesting and novel work with carbon black, but then and even today, many companies are afraid to publish technical work, less they help their competitors. Much of his work is lost in the printed archives at GY.

Following him was Marty Testa who could make the EUM-4 sing. A lot of the morphology work on latex rubber and carbon black mixing at GY was supported by Marty. I still remember the angry polymer chemist who was sure we sabotaged his work when all his latex particles turned out to be oval, or football shaped instead of round.

I understand GY no longer has a TEM, which makes me sad. John Myers and myself might have been their last TEM microscopist.

All things are transitory, I'm depressed and I'm off to look a funed silica...........
Frank

-----Original Message-----
X-from: bigelow-at-umich.edu [mailto:bigelow-at-umich.edu]
Sent: Wednesday, April 11, 2012 11:33 PM
To: Frank Karl

Listers:

I just received the following request from Lee Gould ("Lee Gould"
{lgould-at-nmu.edu} ) and thought it worthwhile forwarding it to you-all:

: " I found a website with RCA microscope information
on it and your name and contact information was provided. I am
looking for some help with RCA EMU-3G and EMU-4. Specifically
Northern Michigan University has one each of these and would like to
get rid of them. I see they could contain Radiation? Any other
hazardous materials? Any one interested in these microscopes? Not
sure you can help, but any information you can provide would be much
appreciated, or any contact information to anyone who can help?
Thank you for your valuable time!"

The RCA EMU-3 was built in the mid 1950s, and the EMU-4 in the mid
1960s, and so both instruments might well be considered "antiques".
I doubt that anyone would be interested in trying to get either back
into operation; however, maybe they would be valuable as a
collector's item or for a museum display.

--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

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This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

==============================Original Headers==============================
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Good Afternoon,
I am a high school science teacher who has been put in charge of learning how to use a recently donated Stereoscan 90. At this time I do not have access to any sample preparation equipment. I was wondering if you may have any suggestions of interesting samples to look at that do not require any preparation for high school students. My students seem to be more interested in looking a biological specimens. Would anyone have samples they may be interested in donating for my students to look at?
Thank you for your time.

Jaimie

==============================Original Headers==============================
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The formula comes from page 66 of "Biomedical Electron Microscopy" by
Maunsbach and Afzelius (http://www.amazon.com/Biomedical-Electron-Microscopy-Illustrated-Interpretations/dp/0124806104/ref=sr_1_1?ie=UTF8&qid=1334265482&sr=8-1
). This is a book that I think very highly of, but have no financial
interest in. This formula has served me well through the years.

David

PS Let the 'out of office' messages begin!

On Apr 11, 2012, at 11:09 PM, Stephane Nizet wrote:

} Hi David!
}
} Interesting calculation! How did you come to this? Empirically?
}
} Regards,
} Stephane
}
} ----- Original Message -----
} From: "dae1-at-email.arizona.edu" {dae1-at-email.arizona.edu}
} To: nizets2-at-yahoo.com
} Cc:
} Sent: Thursday, April 12, 2012 4:37 AM
} Subject: [Microscopy] Re: Fixation for LM
}
}
}
}
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} I use 10% of the Formaldehyde molarity and 30% of the Glut molarity
} and add that to the buffer. This seems to work well for total.
} David
}
}
} On Apr 11, 2012, at 3:42 AM, {nizets2-at-yahoo.com} {nizets2-at-yahoo.com}
} wrote:
}
} }
} }
} }
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} } Dear list,
} }
} } I have a strange behavior during a LM fixation and I would
} } appreciate your feedback.
} } I prepared my fixative as follows:
} } - suspended 2 grams of a 6-years old paraformaldehyde powder in mQ
} } water
} } -Heated at 60�C for 10 min with magnetic stirring.
} } - Added 3 drops of 1N NaOH to depolymerize
} } - let cool down, then double the volume with 100mM Hepes pH 7.2
} } Final solution is 4% PFA in 50mM Hepes pH 7.2
} }
} } I used the solution 2 hours after preparation on cells grown on
} } coverslips. I washed them just once with PBS before adding fixative.
} } Under the LM I can see impressive blebbing of the cell membrane and
} } that's my concern.
} } It seems that my solution is hypoosmotic, is it the case?
} } In this hypothesis I could just add some NaCl to make it isotonic.
} } At the bottom of my memory I seem to remember that isoosmoticity
} } sould be approx. 260mosm.
} } The problem is that I don't know how to take account of the
} } presence of formaldehyde for the calculation of the osmolarity.
} } At the beginnning of the fixation I suppose that only a few
} } molecules cross the membrane but as the fixation proceeds more and
} } more molecules cross the membrane. 4% is quite high so if it counts
} } I guess that the solution is not hypoosmotic.
} }
} } Stephane
} }
} }
} } ==============================Original
} } Headers==============================
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Greetings listserver,

Our research group is seeking access to a full ex-situ lifout station
(for manipulating FIB samples) in the vicinity of Philadelphia, PA.
Please contact me offline to discuss details about the project.

Thank you,
Chris
Drexel University
Philadelphia, PA

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Angel,

I have a copy - I'll have to scan it in and make a pdf of it, though,
so I won't be able to get it to you before next week. If someone
offers a pdf they already have, or a hard copy, please let me know.
You might also want to look into joining the open cut group on gmail
- it's for people like us who have microtomes from the depths of last
century.
joining open cut [1 Update]
{http://groups.google.com/group/open_cut_project/t/64fa2f506e15e235} http://groups.google.com/group/open_cut_project/t/64fa2f506e15e235

Phil

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} Email: Angel.Paredes-at-uth.tmc.edu Name: Angel Paredes
}
} Organization: University of Texas Health Science Center
}
} Title-Subject: [Filtered] Sorvall MT2 instruction manual
}
} Message: Hi,
}
} I just purchased a used Sorvall MT2B ultramicrotome. Would anyone
} have a users manual they can share?
}
} Sincerely,
} Angel Paredes

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Listers,

I have asked Jaimie where she is located so that nearby microscopists may be
able to help her with her previous request (see below).

I am location in Southern California, in Ontario about 30 miles from LA.
Thank you.

Jaimie Graham
Ontario High School
Chemistry
(909) 988-7411 (ext. 2208)
{Jaimie_Graham-at-cjuhsd.net}

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E � Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov

X-from: {Jaimie_Graham-at-cjuhsd.net}
Reply-To: {Jaimie_Graham-at-cjuhsd.net}

Good Afternoon,
I am a high school science teacher who has been put in charge of learning
how to use a recently donated Stereoscan 90. At this time I do not have
access to any sample preparation equipment. I was wondering if you may have
any suggestions of interesting samples to look at that do not require any
preparation for high school students. My students seem to be more
interested in looking a biological specimens. Would anyone have samples
they may be interested in donating for my students to look at?
Thank you for your time.

Jaimie

==============================Original Headers==============================
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-------- Original Message --------

-------- Original Message --------

Jennifer

Look up Moire' fringes. This is an interference effect which
occurs when 2 crystals are misoriented slightly and both are
diffracting. The interference period is a function of both
d-spacings and the misorientation angle between the lattice
directison. Layered materials (clays) can easily do this.
The spacing of the interference fringes can have very large
"apparent" d-spacings.

For a detailed discussion see for example.

L. Reimer Transmission Electron Microscopy.
Chapter 8 page 359- 361 (1989)
Vol 36 Springer Series in Optical Sciences

Your Friendly Neighborhood SysOp
Nestor

On 4/13/12 6:29 PM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Date: Fri, 13 Apr 2012 10:10:59 -0500
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} Email: mille638-at-muohio.edu
} Name: Jennifer Tully
}
} Organization: Miami University/Department of Geology
}
} Title-Subject: [Filtered] TEM micrograph of river sediment fines
}
} Message: I have a micrograph (100kV, 100Kx) of what looks like lattice fringes, however the spacing
} is much too large (~10nm between the lines). The "lattice" is very regular in spacing and perhaps is
} the result of clay layering as the sample contains the fine fraction of river sediment including
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--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory
Fellow of the Microscopy Society of America
E.P. Wigner Fellow - Oak Ridge National Laboratory
Senior Fellow of the Computational Institute - University of Chicago
Past-President Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a Mac !

===========================================

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Morning Marilena,

We have a Tecnai 12T with no accessories beyond a 1k Gatan camera. In 2008(?), or thereabouts, I received a quote for upgrading our instrument to STEM and TOM. There was no discussion of what should be possible since the majority of the upgrade would have been software. The information was useful, but the cost was not affordable or justifiable. The only hardware change would have been the stage IF it was not already capable of the mechanical resolution required for TOM (that replacement was included in the quote since 9 of 10 stages did NOT fall into the usable category.

Our lesson was that the Tecnai series was designed to carry any accessory package manufactured by FEI. The same may be true for you.

Best info - historically: http://www.biochem.mpg.de/doc_tom/Acquisition/tom/Acquisition/tecnai12.html, and
http://www.biochem.mpg.de/doc_tom/Acquisition/tom/Acquisition/FAQ.html

The above TOM software was apparently developed by those folks at www.biochem.mpg.de. Where they may be now I cannot say, but the software was, at one time, available from them.

Hope this helps,

Fred Monson

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

If one looks long enough, one may actually see something frozen in time that flashes one's mind to a new state.
1. The day that I asked myself how the hydrogen atom's mass could actually be: 1.0008.
2. The day, while looking at a slide of some tissue, I saw a leukocyte that had been caught sneaking in or out of a capillary: http://www.annualreviews.org/doi/abs/10.1146/annurev-pathol-011110-130224

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Monday, April 16, 2012 8:51 AM
To: Monson, Frederick

http://www.biochem.mpg.de/baumeister/tom_e/index.html

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

If one looks long enough, one may actually see something frozen in time that flashes one's mind to a new state.
1. The day that I asked myself how the hydrogen atom's mass could actually be: 1.0008.
2. The day, while looking at a slide of some tissue, I saw a leukocyte that had been caught sneaking in or out of a capillary: http://www.annualreviews.org/doi/abs/10.1146/annurev-pathol-011110-130224

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Monday, April 16, 2012 8:51 AM
To: Monson, Frederick

Hello,
to centre the beam on the centre of the screen, just use the SHIFT wheel. It will translate=shift the beam.

If you need to adjust the tilt of the voltage or current axis of the beam to run in parallel to the optical axis, then activate with the "Bright Tilt" button the multifunctional wheel "Def/Stig", and properly adjust=tilt the axis with this. (see manual chapter 5.3.5 for further instructions on this).

To understand what�s happing, please have a look on i.e. this:
http://www.matter.org.uk/tem/scanning_tilting_alignment.htm

You will see that shifting does not only alter the CLA2, but also affects CLA1. They both (CLA1 and CLA2) are acting concerted, in certain relation to each other, on order obtain the translation of the beam.

On your JEM-2100F, the CLA1 effect might appear so small, that you can�t read it out easily from the Lens/Def Voltage Value panel. But if you write down the CLA values with the "Def/Lens Memory" tool before and after manipulating them, then you will see the effect well represented in the changing values.

Best greetings from the EM-Labs of CIC biomaGUNE,
Marco M�ller

----------------------------------------------------------------
Marco M�ller
Platform Manager - Electron Microscopy

CIC biomaGUNE
Parque Tecnol�gico de San Sebasti�n
Paseo Miram�n, 182. Ed. Empresarial C
20009 San Sebasti�n (Guip�zcoa)
SPAIN

Tel:� +34 943 00 53 25
mmoller-at-cicbiomagune.es

-----Original Message-----

Dear Andrea,

for at least an approximate reply/solution of your problem(s) it would be of greatest interest to know about:

1) kind of tissue (assuming, it's biological of nature) (there are some tissues which are prone to wrinkling after staining)
2) at least some data about your tissue processing
3) {age} of embedding kit (e.g. also how often the bottle/kit was used before, catalyst/hardener which one, how old is this)
4) composition of your embedding resin (incl. catalyst/hardener), soft-hard mixture
5) composition of your staining solution (hydrous, alcoholic, percentage) assuming you stain as usual/classically UO2Ac followed by Lead (citrate).
6) also it would be of interest in which way you are placing your grids with the sections in(to) the staining solutions (using automated staining device, a special staining device for manual staining, or just a quadratic glass through with a well ['embryo dish'] when staining manually, the latter procedure perhaps prone to such artifacts as you describe...tweezers, drying the sections - grid within tweezers - and so on...
Perhaps you'll inform us...

best wishes and regards,

Wolfgang MUSS, PhD
EM-Lab
SALK-LKH/Pathology
Salzburg, Austria

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} Gesendet: Samstag, 14. April 2012 01:32
} An: Mu� Wolfgang
} Betreff: [Microscopy] Embed 812 wrinkled thin sections (UDS after staining==} wrinkles)
}
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} Email: kaszas.1-at-osu.edu
} Name: Andrea Kaszas
} Organization: Ohio State University/OARDC
} Title-Subject: Embed 812 wrinkled thin sections
} Message: Dear listeners,
}
} I use Embed 812 embedding kit and I section 70 nm thickness sections
} for TEM study.
} My sections are fine until staining, no wrinkles.
} After staining I have wrinkles on my sections.
} Any suggestion would be greatly appreciated.
} Thanks for your time in advance.
}
} Andrea Kaszas
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Hi

Both the gun and the complete illuminating system, gun plus condenser
lenses, need to be centered and tilted to place the electron beam absolutely
straight down the magnetic axis of the system.

In the case of the gun, it is "shifted" onto the screen centre and the angle
of the beam in relation to the magnetic axis of the first condenser lens is
adjusted by the gun "tilt" controls. In order to make this alignment the
second condenser is brought to crossover, imaging the virtual source. This
action is best carried out at a magnification that makes the image about
4cms across. The gun shift must be used to centre the beam on the screen
whilst the gun tilt is used to evenly balance the halo of the desaturated
source.

In the case of the illuminating system, once the gun is aligned as described
the complete illuminating system is shifted to centre the beam on the screen
and tilted to place the beam on the magnetic axis of the objective lens.
The latter action dependes upon the microscope. Japanese instruments allow
this alignment to be carried out either to the current centre of the
objective lens or to the voltage centre. European instruments simply align
the current centre. For the current alignment the objective lens current is
changed and the illumination tilt used to centre the spinning pulse on the
screen. In the voltage alignment case the high voltage level is changed and
the illumination tilt is used to centre the image pulse on the screen. The
changing the illumination tilt controls will change the centre of the
magnification/diffraction pattern on the screen thus further "projector"
alignment may be required.

When aligning the instrument it is very importenat when working on the gun
to ONLY align the instrument with the GUN controls; a big error made by
many! Under normal operating conditions the illumination should be kept in
alignment with the illumination shift controls. This action is eccential
because the shift has no influence on the voltage or current alignments
which are very critical alignments.

Enjoy

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
www.emcourses.com

.} Email: amit.welcomes.u-at-gmail.com
} Name: Amit
}
} Organization: Indian institute of science
}
} Title-Subject: [Filtered] difference between "Shift" and "Bright Tilt"
}
} Message: While using Jeol 2100f i noticed one thing that the "Bright Tilt"
} button on left panel
} alters one of the deflector coils above condenser lens( CLA1). while the
} beam "shift" knob alters
} the other deflector coils (CLA2). why is one "Shift" then and other one is
} "Tilt"? is there any
} logical explanation on when to use which one for bringing beam to centre?
}
} Thank you
}
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Dear Listers:

I would like to bring your attention the following events which may be of your interest:

1-For those who are interested to explore Rhodes Island/Greece, there will be Advanced Characterization session at the International Conference on Nanostructured Materials �NANO2012� to be held in Rhodes Island, Greece. The session will highlight in-situ and ex-situ characterization techniques of nanostructured materials. Papers in this conference will be reviewed by members of the scientific committee in order to be published as a special issue of a relevant scientific journal. For more information, please visit www.nano2012.org

2-The Journal of Nanotechnology will have a Special Issue on Characterization of Energy-Related Materials. For important dates and topics, please visit www.hindawi.com/journals/jnt/si/502851/cfp/

Best Regards,

Sincerely,

Jafar

Jafar F. Al-Sharab, Ph.D.

Rutgers, The State University of New Jersey
Department of Materials Science and Engineering
McLaren Center for Ceramic Research
607 Taylor Road, Room 237
Piscataway, NJ 08854-8065
Tel. 848-445-5580
Fax 732-445-3258

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Hi Michael!
�
DRAQ5 intercalates in DNA, it is not necessarily the same kind of (strong) interactions as the one between antigen/antibodies.
So it probably dissociates with time.
I see 3 solutions:
The easiest one is probably to store your samples at -80�C or -20�C. Easy but�the storage volume in freezers is not unlimited. Morevoer, first make sure that freezing/thawing does not affect your staining, since it could very well disrupt cellular structures important for you.
Another solution would be perhaps to re-incubate with DRAQ5 (it costs money).
Alternatively you could try to fix your sample after the whole staining procedure. However there is no guarantee that the fixation stabilizes DRAQ5-DNA bound.
�
Have fun with 3-color stainings.
�
Stephane

________________________________
X-from: "microscopylistserver-noreply-at-microscopy.com" {microscopylistserver-noreply-at-microscopy.com}
To: nizets2-at-yahoo.com
Sent: Saturday, April 14, 2012 1:32 AM

-------- Original Message --------

Good afternoon,
Dear Juan Luis,

it seems to me (if I remember correctly) that 'yeast' ...."is the beast" and a challenging task in {classical} preparation for TEM [i.e. chem. fixation, dehydration, embedding in resins]...
Most papers published obviously show these problems in optimal ultrastructural preservation.

A lot of papers recommend freezing and cryosectioning, Freeze-substitution, embedding in e.g. Lowicryl(s) or LR White with / without / by means of the PLT-(progressive lowering temperature-)Method etc.

Not having the possibility to do HPF in our lab but knowing about some papers published on this topic,
e.g. cf.: www.formatex.info/microscopy4/11-18.pdf

Investigation of the ultrastructure of yeast using cryo-immuno-electron microscopy
Muriel Mari, Janice Griffith and Fulvio Reggiori
in: Microscopy: Science, Technology, Applications and Education
A. M�ndez-Vilas and J. D�az (Eds.) pp.11-18; � FORMATEX 2010 with an interesting new IEM method (rehydration method, permitting optimal preservation of the epitopes)
" Fixation and subsequent embedding in resins such as Spurr's and Epon, has allowed for obtaining satisfactory results in preserving the yeast morphology. With conventional EM approaches, the more prominent subcellular compartments such as the nucleus or the vacuoles are readily detectable while small organelles, including the different Golgi compartments or the various classes of endosomes, cannot be easily seen." and:
"One of the major challenges in the processing of yeast for EM investigations has been the relative impermeability of the cell to both fixatives and resins due to the presence of a cell wall."

an older publication of the same authors:
A Cryosectioning Procedure for the Ultrastructural Analysis and the Immunogold Labelling of Yeast Saccharomyces cerevisiae
Janice Griffith, Muriel Mari, Ann De Mazie`re and Fulvio Reggiori
Traffic 2008; 9: 1060-1072 (Blackwell Munksgaard) you can find on:
http://onlinelibrary.wiley.com/doi/10.1111/j.1600-0854.2008.00753.x/abstract
http://onlinelibrary.wiley.com/doi/10.1111/j.1600-0854.2008.00753.x/pdf

An older thread previously from (MSA!) listserver (5/8/1997) concerning the fixation / embedding problem you can find on e.g.: http://www.biotech.ufl.edu/EM/data/potatotem.html

Another trial worth perhaps would be "microwave assisted fixation and processing": e.g.
see: http://attach.sciencedirect.com/science/article/pii/S1434461011000484
Protist, Volume 163, Issue 2, March 2012 , Pages 296-305
Improvement of Ultrastructural Preservation of Eimeria Oocysts by Microwave-assisted Chemical Fixation or by High Pressure Freezing and Freeze Substitution
Thomas Kurth, Stefanie Wiedmer, Rolf Entzeroth
{ {Fixation and preparation for electron microscopy of coccidian oocysts is a general problem. Especially in sporulated oocysts, proper fixation and resin infiltration are hindered by the robust oocyst wall. Conventional chemical fixation therefore leads to artefacts that obscure cellular details in the oocysts.

In this study, sporulated oocysts of Eimeria nieschulzi were subjected to different fixation and embedding protocols: conventional chemical fixation and embedding in Spurr's resin, microwave-assisted fixation and processing followed by embedding in epon, and high pressure freezing followed by freeze substitution and epon embedding. The samples were finally studied by transmission electron microscopy. Many ultrastructural features of the oocyst wall and the sporozoites were already substantially improved after microwaved-assisted fixation and processing. However, the fine structural preservation still suffered from shrinkage and artificial extraction, which occured during dehydration and infiltration. High pressure freezing (HPF) and freeze substitution (FS) revealed much better preservation. Oocyst walls retained their ovoid shape, and the ultrastructure of sporozoites was well preserved with no signs of shrinkage or extraction. HPF and FS are therefore a suitable method for the ultrastructural analysis of coccidian oocysts
Keywords: Eimeria; electron microscopy; freeze substitution; high pressure freezing; microwave cell processing; oocysts } } unfortunately tat paper isn't free of cost, but Author's affiliation and e-mail for requesting the paper would be:
Thomas Kurth, TU Dresden, DFG-Center for Regenerative Therapies Dresden, Tatzberg 47/49, D-01307 Dresden, Germany thomas.kurth-at-crt-dresden.de

So, at the end here, unfortunately I can't provide a 110% optimal recipe but perhaps - what I wrote - is helpful anyway.

Best wishes and regards,
greetings to Sevilla / Seville (I am going to visit your beautiful city in September this year (:-)) )

Wolfgang MUSS, PhD
EM-Lab, Pathology
SALK-LKH (Gen. Hospital)
SALZBURG - AUSTRIA

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} Gesendet: Dienstag, 17. April 2012 14:41
} An: Mu� Wolfgang
} Betreff: [Microscopy] Yeast staining protocol
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} Email: jlribas-at-us.es Name: Juan Luis Ribas
} Organization: Microscopy Facility. University of Seville. Spain
} Title-Subject: Yeast staining protocol
} Message:
}
} Good morning,
}
} We are trying to work with yeasts in Spurr and we are having poor
} sample preservation. We have tried
} to increase the incubation times, the permanganate alternative and the
} partial digestion of the cell
} wall but the protocols are not working fine. Would it be possible to
} share your experiences with
} these sample protocols?
} Best regards
}
} --
} Juan Luis Ribas
} Servicio de Microscop�a
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Hi Nancy,
I don't know that this particular company has been talked about previously,
but the general topic of insurance companies covering equipment maintenance
has been covered many times in the archives.

If memory serves me correctly, the overwhelming response is "Don't do it!"
with a few exceptions of people who have been satisfied.

Most complaints center around:

1) Bickering over whether or not some particular issue is covered and
2) Suddenly becoming a billable customer in the manufacturer's eyes and
going to the bottom of the list in terms of response time to a problem.

Search the microscopy archives over the past few years and I think you'll
find a number of extensive threads.

Disclaimer: Quality Images provides actual service contracts for SEMs in
the northeastern US.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
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kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com
[mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Wednesday, April 18, 2012 9:04 AM
To: kenconverse-at-qualityimages.biz

Hi All,

I had an inquiry about using the following fix for mice perfusion with the
intent to use tissues for TEM. I had never seen it used for general
fixation so wondered if anyone had experience with it.

1.5% glutaraldehyde, 4% polyvinylpyrrolidone, 0.05% calcium chloride, and
0.1 M sodium cacodylate, pH 7.4.

Debby

--
Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy/

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I used a similar fix many years ago to fix the renal medulla of the rat,
gives good preservation of the intercellular spaces by increasing the
colloid osmotic pressure of the fix.
I think Maunsbach introduced PVP in the early 1970's. Try J. Ultrastruc.
Res. 30:195, 1970.
1.5% glut sounds "low" but is plenty.

Geoff

On 4/18/2012 12:45 PM, dsherman-at-purdue.edu wrote:
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} Hi All,
}
} I had an inquiry about using the following fix for mice perfusion with the
} intent to use tissues for TEM. I had never seen it used for general
} fixation so wondered if anyone had experience with it.
}
} 1.5% glutaraldehyde, 4% polyvinylpyrrolidone, 0.05% calcium chloride, and
} 0.1 M sodium cacodylate, pH 7.4.
}
} Debby
}

--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583
mcauliff-at-umdnj.edu
**********************************************

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Dear Debby,
apologize for not knowing whether you got a lot of messages or not off List or not,
hoping that my post-reply (yet lengthy) is of any help.

Regarding the use of PVP I remember from my "really early, ancient" times (exptl. studies for my thesis, when I had to perfusion-fix rat brains for resin-embedding, serial sectioning and 3Dim-reconstruction) that I used this {chemical} ('filler substance')
a) with the / a 'physiological' Tyrode buffer ('mammal' ringer solution) for flushing the circulation system and
b) with the fixative (perfusion fixation, by means of a self made perfusion apparatus, capable of varying perfusion pressure for smaller and bigger animals, according to the un-/known blood pressure of the animals circulation system).

I also remember that there were several "kinds" of PVP (different mol. weights) and if I am correct, I used PVP-40 (average mol. weight 40,000, K-value 28-32) from SIGMA. I don't remember now the percentage (this I would have to check with old protocols not at hand for now.
I have seen in the specific literature that PVP-40 is used often, but the PVP {xx Mol. weight} used might depend on the animal you will "treat" as well as the organ system you'll need to have fixed optimally (see the literature, mostly in the 70ies, rare exceptions up to 2010, some saved in file at least as lit. ref's).

Ok, chewing over, I chased after and indeed found some records now, which I wrote down in 1987-1988, when I was asked to (perfusion-) fix PIG kidneys (mini-pigs, exptl. surgery) for TEM.
Hoping not to bother you or any Lister here, I honestly would like to place here some of those notices
(mOsmol measurements always done 3 times with another pipetted aliquot):

1) (plain) Tyrode solution (after HAYAT 1981, p 411, + 1 g Glucose): should measure 290 mOsmol (measured values:
pH approx. 7.5, but once also pH 8.1 due to longer storage; 278, 282, 284, 284 mOsmol- so one should add glucose perhaps too)
2) as 1) freshly prepared, 4 degr. C, plus 2.5 PVP-40 (rigorous and longer { at least 10 min} mixing/stirring necessary!): pH 7.5-7.6; 293-294, 298-299, 295-296 mOsmol
3) 25 ml GA (25%)[end concentration in fixative: 0.625 % GA] + approx.900 ml Tyrode solution as in 1) + 1 g glucose ad 1000 ml end volume with Tyrode(as in 1)): pH=7.6; 358-359; 364-365; 363-364 mOsmol.
4) 2.5%(? I am not quite sure here... am not really powerful in such calculations) GA-Tyrode solution (= 5 ml 25% GA ad 50 ml Tyrode) + 2.5 % PVP-40: pH 7.5-7.6; 647-648; 641-642; 655-656 mOsmol.

X-from all my records it seems that longer storage at RT or 4 degr.C ends up with an increase of pH (e.g. after 7-8 hrs: pH 8.3 - 8.35; 8.45-8.5).

The TYRODE-Ringer solution which is mention in MILLONIG G.(iusepppe)'s "Laboratory Manual for Biological Electron Microscopy (1976) [NB: very informative "methods" booklet!] consists of:
STOCK solutions:
Solution A (pH approx. 4.8) :
NaCl 20.00 g
KCl 00.50 g
CaCl2.6H2O 00.50 g
MgCl2.6H2O 00.25 g
H2O to make 100.00 ml

Solution B(pH approx. 8.5 [measured: 8.1-8.4]):
NaHCO3 05.00 g
NaH2PO4 00.25 g
H2O to make 100.00 ml

Working Solution: Mix (40ml soltn A + 460 ml H2O) + (20 ml soltn B + 480 ml H2O) + add 1.0 g Glucose
This solution / recipe has been overtaken by M.A. HAYAT 1981, p. 411
Own measurement:
(40ml soltn A + 460 ml H2O) resulted in pH 5.95
(20 ml soltn B + 480 ml H2O) resulted in pH 9.1
mixed together: pH= 8.6-8.7 so you perhaps have to calibrate with 1 N HCl (+ 1 ml / 1000 ml) resulted in pH 7.5.

There also is listed (in Millonig's Manual) a
Mammalian Ringer(MamR), which perhaps suits better for mice or rats, but don't know /have no experience with this:
MamR (1):
H2O to make 200.00 ml
NaCl 1,80 g
KCl 0.84 g
CaCl2.6H2O (CaCl2.2H2O) 0.48 g (0.40 g)
NaHCO3 0.10 g
Glucose 0.10 g
MgCl2.6 H2O 0.05 g

MamR(2)
H2O 200.00 ml
NaCl 1.80 g
KCl 0.84 g
CaCl2.6H2O 0.36 g

(all components to be solved one after the other, to prevent precipitation in the solution)

MamR (1) measured: pH 7.7; 446;446; 447 mOsmol; after 50 hrs: pH 8.15; 448-449; 450-452; 449-450 mOsmol
(NB: measurements are my own values, and are not included in the Millonig's Lab. Manual, respectively)

If you add PVP-40 (e.g. 2.5%) you should not measure a very different mOsmol-value with an osmometer, I think..

For optimal preservation of ultrastructure -though- one has not only to consider the solutions but - more stringent - the correct perfusion pressure, temperature (room temp. considered to be better than 4 degr. C) and flow volume/time through the chosen artery, optimal flushing pre-perfusion to get rid of blood cells.
Flushing of the circulation system should be finished at least after 2-4 mins (no more blood e.g. from V. cava). Perfusion fixation should be finished after some 5-10 min -at- RT, then dissect chosen organ systems and postfix with the fixative [without PVP added) for another 1-3 h -at- RT.
(Procedure of preparation - not with a PVP-40 containing fixative) cf. e.g.
http://www.google.at/url?sa=t&rct=j&q=perfusion%20fixation%20and%20vena%20cava&source=web&cd=4&ved=0CEkQFjAD&url=http%3A%2F%2Fwww.gladstone.ucsf.edu%2Fgladstone%2Ffiles%2Fhistology%2FProtocols%2Ffixingperkouse.pdf&ei=_TuQT-euPIPKsgba47iKBA&usg=AFQjCNGEGXKR944yL0wWL3VkIyMHzri3xA)
Sorry, this pdf you'll get perhaps only with that long URL (==Percent-encoding, also known as URL encoding )
www.gladstone.ucsf.edu/.../fixingperkouse.pdf� since:
www.gladstone.ucsf.edu/gladstone/files/histology/Protocols./fixingperkouse.pdf will not work! (pdf saved in my files).
�

More measurements of solutions on request. Specific Literature (at least References) also in file (so if interested send an e-mail requesting these)

I shall end up here with greetings to you and the listers who perhaps are looking also for recipes with this.

Best regards and wishes,

Wolfgang MUSS
EM-Lab, Pathology
SALK-LKH (Gen. Hosp.)& PMU Salzburg
SALZBURG-Austria

Von: dsherman-at-purdue.edu [mailto:dsherman-at-purdue.edu]
Gesendet: Mittwoch, 18. April 2012 18:48
An: Mu� Wolfgang
Betreff: [Microscopy] Polyvinylpyrrolidone use
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Hi All,

I had an inquiry about using the following fix for mice perfusion with the intent to use tissues for TEM.
I had never seen it used for general fixation so wondered if anyone had experience with it.

1.5% glutaraldehyde, 4% polyvinylpyrrolidone, 0.05% calcium chloride, and 0.1 M sodium cacodylate, pH 7.4.

Debby

--
Debby Sherman, Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy/

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Has anyone had a problem with magnetic beads causing charging artifact in the SEM?

We're operating a Hitachi S2600H using secondary electron detection (acceleration voltage=20kV; emission current=100uAmp; beam current=20; WD=15mm). The biological samples (mouse glomeruli isolated using Invitrogen Dynabeads--still present) were fixed onto poly-L-lysine-coated cover slips, osmicated, dehydrated, chemically dried with HMDS and sputter-coated with AuPd. (I checked for obvious causes--the stubs were snug, etc.)

Thank you now (and for all your help in the past),

Jaclynn Lett

Senior Research Technician, EM Facility
Research Center for Auditory and Vestibular Studies
Department of Otolaryngology
Washington University School of Medicine
660 S Euclid Ave, Campus Box 8115
St. Louis, MO 63110

Office: 314-747-7257
Fax: 314-747-7230
Email: lettj-at-ent.wustl.edu
http://otocore.wustl.edu

The materials in this email are private and may contain Protected Health Information. If you are not the intended recipient, be advised that any unauthorized use, disclosure, copying, distribution or the taking of any action in reliance on the contents of this information is strictly prohibited. If you have received this email in error, please immediately notify the sender via telephone or return email.

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-------- Original Message --------

-------- Original Message --------

Hi

Double check the path to earth, that the sample is in good electrical
contact with the stub as a first step. However for the type of specimen that
you are using, even though you have sputter coated, 20kV is far too high.
The only reason I would use 20kV on a specimen like this is if I was after
sub surface detail in backscatter. Similarly the emission current is too
high for this type of specimen; too many electrons for the path to earth to
carry!

Sputter coating is not always the total answer to charge. Accelerating
voltage relates to penetration and the greater the penetration on some
samples the greater the charge level even when coated. Look at dropping to
10kV with an emission current of around 50 to 75uA and keep the spot
size/probe current well down. You do have the best working distance (15)
for low charge but the other parameters are causing you the problems I feel.

Good luck

Steve

Steve Chapman
Protrain for Consultancy and Courses in Electron Microscopy
www.emcourses.com

-----Original Message-----
X-from: LettJ-at-ent.wustl.edu [mailto:LettJ-at-ent.wustl.edu]
Sent: 19 April 2012 17:46
To: protrain-at-emcourses.com

Has anyone had a problem with magnetic beads causing charging artifact in
the SEM?

We're operating a Hitachi S2600H using secondary electron detection
(acceleration voltage=20kV; emission current=100uAmp; beam current=20;
WD=15mm). The biological samples (mouse glomeruli isolated using Invitrogen
Dynabeads--still present) were fixed onto poly-L-lysine-coated cover slips,
osmicated, dehydrated, chemically dried with HMDS and sputter-coated with
AuPd. (I checked for obvious causes--the stubs were snug, etc.)

Thank you now (and for all your help in the past),

Jaclynn Lett

Senior Research Technician, EM Facility
Research Center for Auditory and Vestibular Studies
Department of Otolaryngology
Washington University School of Medicine
660 S Euclid Ave, Campus Box 8115
St. Louis, MO 63110

Office: 314-747-7257
Fax: 314-747-7230
Email: lettj-at-ent.wustl.edu
http://otocore.wustl.edu

The materials in this email are private and may contain Protected Health
Information. If you are not the intended recipient, be advised that any
unauthorized use, disclosure, copying, distribution or the taking of any
action in reliance on the contents of this information is strictly
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notify the sender via telephone or return email.

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} Reply-To: Lee Cohen-Gould {lcgould-at-med.cornell.edu}
} Below is the result of your form, submitted on Friday, April 20,
} 2012 at 09:10:50 AM.
}
} realname - Lee Cohen-Gould
} Email - lcgould-at-med.cornell.edu
} EDUCATION - Graduate College
} LOCATION - USA
} SUBJECT_OF_QUESTION - multiple labels in immunogold
} QUESTION - Hi All-
} Nestor's filters keep rejecting me, so I'm coming in the back door.
} I have a client who hopes to do triple immunogold labeling. I have
} been noodling around with ideas including peroxidase-DAB or
} peroxidase-ENZMET for one of the labels and 2 sizes of gold for the
} others. I thought that I'd toss this out as an intellectual
} exercise to those of you who do this all the time. Here's another
} complication: she has 3 primary antibodies and all are available
} from rabbit or goat hosts, but that's it. Two of the primaries will
} have to be from the same host. I thought that I'd try Protein A or
} G for one of them, using excess, untagged protein to block before
} the second primary is used. Long and tedious. Any better ideas?
} I know I can count on THE LIST for suggestions.
} Thanks in advance!
} Lee

==============================Original Headers==============================
3, 24 -- From oshel1pe-at-cmich.edu Fri Apr 20 11:13:59 2012
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Check out the article below.

An Innovative Triple Immunogold Labeling Method to Investigate the
Hemopoietic Stem Cell Niche In Situ

Sarah L. Ellis, Brenda William, Stephen Asquith, Ivan Bertoncello and
Susan K. Nilsson

Microscopy and Microanalysis (2009), 15 : pp 403-414

Hong

On 4/20/12 12:16 PM, "oshel1pe-at-cmich.edu" {oshel1pe-at-cmich.edu} wrote:

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Dear All, our lab is intending to purchase a plasma cleaner for the TEM sample holder. Right now I see the available ones from Gatan (Solarus Model 950)and Fishione (Model 1020 and 1070). May I ask if any of you use these models and able to comment on it? On the other hand, are there any�advises�that you can give in the features that would be good to have for a Plasma Cleaner? I am also glad if there are some other brands that you have used before.

Thanks in advances! Have a great weekend!

Cheers,
Yee Yan, Tay
FACTS Lab
NTU

==============================Original Headers==============================
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4, 35 -- Subject: Need Advises for Plasma Cleaner for TEM Sample Holders
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Hi Yee Yan
you might want to compare the new Hitachi Zone Cleaner against a
plasma cleaner before you decide.
Elaine

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--

Dr. Elaine C. Humphrey
STEHM Technologist and Lab Manager
Bob Wright Science Centre A015
Advanced Microscopy Facility
University of Victoria, Canada

Lab: 250-853-3968
cell: 250-886-2068
website: http://www.stehm.uvic.ca

==============================Original Headers==============================
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Dear Yee Yan,
besides the well-known manufacturers there is a plasma-cleaner available
from a German producer which is very good (just a satisfied customer...)
and often used at German institutes...
Please see http://www.binder-labortechnik.de/ and click on
"plasmacleaner". There is also an English description and manual available.

Best regards,
Stefan

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

Am 21.04.2012 18:57, schrieb one_twinklestar-at-yahoo.com.sg:
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} Dear All, our lab is intending to purchase a plasma cleaner for the TEM sample holder. Right now I see the available ones from Gatan (Solarus Model 950)and Fishione (Model 1020 and 1070). May I ask if any of you use these models and able to comment on it? On the other hand, are there any advises that you can give in the features that would be good to have for a Plasma Cleaner? I am also glad if there are some other brands that you have used before.
}
} Thanks in advances! Have a great weekend!
}
} Cheers,
} Yee Yan, Tay
} FACTS Lab
} NTU
}
}
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} 4, 35 -- From: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
} 4, 35 -- Reply-To: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
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Yee Yan

Plasma "cleaners" are sometimes also called plasma "ashers".

They are commerically manufactured by

South Bay Technology
XEI Scientific
Structure Probe Inc.
Fischione Instruments
Gatan Inc.
Ibis Group
Binker-labortechnik
FEI (has a version which is built into some of their columns)

There is at least one other commerical organization who manufactures
a model but at the moment I cannot remember their company moniker
(my apologies to that company).

In addition to a plasma you can use UV irradiation with a vacuum
assist to also mitigate/remove mobile hydrocarbons.
Of course a light very very low energy ion milling can also be used,
but the UV/Plasma can be adjusted to much lower effective energies.

In principle the technology is fairly straightforward. The key features to
compare between systems are:

Gas composition
Gas pressure control
Plasma power control
Cleaning time control
SEM vs TEM configurations
as well as ease of use.

There are pro/con aspects to each of the above points. Read
each manufacturer's literature closely. Look for examples they
have of applications to material systems similar to those you will
be studying ( i.e. semiconductors, metals, films, etc....) Each
manufacturer will have preset or standard conditions which work best
in their respective geometries, and they are all slightly different.

The best thing to do is to visit someone that has one
of the systems you are considering and actually test it
using the type of specimen you will be studying.

Remember the prime target of these devices is removal of
mobile or surface bound hydrocarbons. So if your "specimen" contains
carbon then in addition to cleaning off any mobile species might
migrate to the electron beam area, it will also sputter away your
sample. So be careful if your planning to use this
for carbon nanotube work!

Disclaimer: My employer Argonne National Laboratory
holds the original patent on plasma cleaning technology for use
in EM applications. A number of the above companies are licensee's
of that technology.

Cheers,

Nestor
Your Friendly Neighborhood SysOp

---===[|]===---

--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory
Fellow of the Microscopy Society of America
E.P. Wigner Fellow - Oak Ridge National Laboratory
Senior Fellow of the Computational Institute - University of Chicago
Past-President Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a Mac !

===========================================

==============================Original Headers==============================
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Nestor humbly neglected to say that if you look up the patent that his
employer holds, Nestor is listed as the Inventor.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: zaluzec-at-aaem.amc.anl.gov [mailto:zaluzec-at-aaem.amc.anl.gov]
Sent: Saturday, April 21, 2012 3:25 PM
To: s.walck-at-cox.net

Yee Yan

Plasma "cleaners" are sometimes also called plasma "ashers".

They are commerically manufactured by

South Bay Technology
XEI Scientific
Structure Probe Inc.
Fischione Instruments
Gatan Inc.
Ibis Group
Binker-labortechnik
FEI (has a version which is built into some of their columns)

There is at least one other commerical organization who manufactures a model
but at the moment I cannot remember their company moniker (my apologies to
that company).

In addition to a plasma you can use UV irradiation with a vacuum assist to
also mitigate/remove mobile hydrocarbons.
Of course a light very very low energy ion milling can also be used, but the
UV/Plasma can be adjusted to much lower effective energies.

In principle the technology is fairly straightforward. The key features to
compare between systems are:

Gas composition
Gas pressure control
Plasma power control
Cleaning time control
SEM vs TEM configurations
as well as ease of use.

There are pro/con aspects to each of the above points. Read each
manufacturer's literature closely. Look for examples they have of
applications to material systems similar to those you will be studying (
i.e. semiconductors, metals, films, etc....) Each manufacturer will have
preset or standard conditions which work best in their respective
geometries, and they are all slightly different.

The best thing to do is to visit someone that has one of the systems you are
considering and actually test it using the type of specimen you will be
studying.

Remember the prime target of these devices is removal of mobile or surface
bound hydrocarbons. So if your "specimen" contains carbon then in addition
to cleaning off any mobile species might migrate to the electron beam area,
it will also sputter away your sample. So be careful if your planning to
use this for carbon nanotube work!

Disclaimer: My employer Argonne National Laboratory holds the original
patent on plasma cleaning technology for use in EM applications. A number
of the above companies are licensee's of that technology.

Cheers,

Nestor
Your Friendly Neighborhood SysOp

---===[|]===---

--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory Fellow of the Microscopy
Society of America E.P. Wigner Fellow - Oak Ridge National Laboratory Senior
Fellow of the Computational Institute - University of Chicago Past-President
Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a Mac !

===========================================

==============================Original Headers==============================
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Dear all,

} Binker-labortechnik
this company (based in Germany) is named "Binder Labortechnik" (not Binker).

} There is at least one other commerical organization who manufactures
} a model but at the moment I cannot remember their company moniker
} (my apologies to that company).

Nestor, was it Harrick Scientific? I am using a Harrick Plasma Cleaner / Sterilizer since } 20yrs now, with great satisfaction (no; I am only a satisfied customer, not a share holder or ...) -- I don't know if they still manufacture their plasma cleaner, though. - I am sure that all the other machines do the same job, basically.
There is nothing else to add, from my point of view. One point, from a practical point of view: for me as a biologist, it is THE method to render carbon support films, grids, etc hydrophilic. Great, and works; always, and no con's (yes, you only have to buy the machine).
kind regards,
Reinhard

--
Prof. Dr. Reinhard Rachel
University of Regensburg
Centre for EM / Anatomy
Faculty of Biology & Preclin. Med.
Universitaetsstrasse 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-regensburg.de
office: VKL 3.1.29

next microscopy conferences:
http://www.emc2012.org.uk/
EMC 2012 in Manchester, UK
http://www.mc2013.de/
MC2013 in Regensburg, Germany

==============================Original Headers==============================
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Hello Yee Yan,
from the conference in Kiel last year, I remember a company called FIB,
which offers a plasma asher
(downstream asher) to clean all TEM, and SEM columns including the samples
inside
or the gentle asher to clean the samples before inserting them.
http://fib.technologie.officelive.com/GV10x.aspx
But I have no experience with them.

Best regards!

Roland

==============================Original Headers==============================
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Dear Dr Nestor and the rest! Thank you very much for your kind advises! Before i ask any further questions, let me do a more in depth studies for the points highlighted!!!

Do have a great day ahead!!!

Cheers,
Yee Yan, Tay
FACTS Lab
NTU, Singapore

----- Original Message -----
X-from: "zaluzec-at-aaem.amc.anl.gov" {zaluzec-at-aaem.amc.anl.gov}
To: one_twinklestar-at-yahoo.com.sg
Cc:
Sent: Sunday, 22 April 2012, 6:15

Yee Yan

Plasma "cleaners" are sometimes also called plasma "ashers".

They are commerically manufactured by

South Bay Technology
XEI Scientific
Structure Probe Inc.
Fischione Instruments
Gatan Inc.
Ibis Group
Binker-labortechnik
FEI (has a version which is built into some of their columns)

There is at least one other commerical organization who manufactures
a model but at the moment I cannot remember their company moniker
(my apologies to that company).

In addition to a plasma you can use� UV irradiation with a vacuum
assist� to also mitigate/remove� mobile hydrocarbons.
Of course a light very very low energy ion milling can also be used,
but the UV/Plasma can be adjusted to much� lower effective energies.

In principle the technology is fairly straightforward.� The key features to
compare between systems are:

�� Gas composition
�� Gas pressure control
�� Plasma power control
�� Cleaning time control
�� SEM vs TEM configurations
�� as well as ease of use.

There are pro/con aspects to each of the above points. Read
each manufacturer's literature closely. Look for examples they
have of applications to material systems similar to those you will
be studying ( i.e. semiconductors, metals, films,� etc....) Each
manufacturer will have preset or standard conditions which work best
in their respective geometries, and they are all slightly different.

The best thing to do is to visit someone that has one
of the systems you are considering and actually test it
using the type of specimen you will be studying.

Remember the prime target of these devices is removal of
mobile or surface bound hydrocarbons.� So if your "specimen" contains
carbon then in addition to cleaning off any mobile species might
migrate to the electron beam area, it will also sputter away your
sample.� So be careful� if your planning to use this
for carbon nanotube work!

Disclaimer:� My employer Argonne National Laboratory
holds the original patent on plasma cleaning technology for use
in EM applications.� A number of the above companies are licensee's
of that technology.

Cheers,

Nestor
Your Friendly Neighborhood SysOp

� � � �� ---===[|]===---

--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory
Fellow of the Microscopy Society of America
E.P. Wigner Fellow - Oak Ridge National Laboratory
Senior Fellow of the Computational Institute - University of Chicago
Past-President Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a� Mac !

===========================================

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Hi all

I remember someone gave once a trick to clean away dried colloidal
silica from a polished sample. I've made a search in the archives, but
I didn't find the post.
So the question is open again.
OK, I know, it's better to rince abundantly immidiatly to avoid the
problem, then to get a solution when the damage is done ! ;-)

Thanks

Jacques

--

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Mat�riaux de Strasbourg
D�partement Surfaces et Interfaces
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail : Jacques.Faerber-at-ipcms.unistra.fr

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Hi Yee Yan,

ibss Group, Inc. exhibited the GV10x DS Asher and Gentle Asher Chamber in
Kiel www.mc2011.de as Roland mentions. At MC 2011 ibss Group, Inc. was
represented by www.fib-kassel.de our German representative.

Visit our web site to view the flexibility offered by the GV10x to clean
specimens on the bench and remove hydrocarbons from chamber and on specimen
at same time.

Please contact us directly for more information.

Best regards,

Vincent Carlino

Disclaimer: ibss Group, Inc. manufacturers both GA10x DS Asher and
GentleAsher Chamber.

ibss Group, Inc.
+1.415.566.5774
+1.415.566.9779 fax
vince.carlino-at-ibssgroup.com
www.ibssgroup.com

-----Original Message-----
X-from: roland.schierholz-at-icmse.csic.es
[mailto:roland.schierholz-at-icmse.csic.es]
Sent: Monday, April 23, 2012 7:25 AM
To: vcrvince-at-comcast.net

Hello Yee Yan,
from the conference in Kiel last year, I remember a company called FIB,
which offers a plasma asher (downstream asher) to clean all TEM, and SEM
columns including the samples inside or the gentle asher to clean the
samples before inserting them.
http://fib.technologie.officelive.com/GV10x.aspx
But I have no experience with them.

Best regards!

Roland

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My Corning stir hotplates that I use for drying thick sections onto glass slides have seen better days.
And there is next to no control much less any temperature measurement.
We just turn them on, barely, then let the heat up. We test the heat by touching the hot plate.
If you can count to 5 before you have to remove your finger, then the plate is hot but not too hot.
Does anyone have a hotplate that they would recommend I consider?
Thanks for you help in advance
Bob
Dr. Robert J. Schmitz
Associate Professor of Anatomical Sciences
Department of Biology
University of Wisconsin-Stevens Point
800 Reserve Street, 380 TNR Bld
Stevens Point, WI 54481
715-346-2420
Email: rschmitz-at-uwsp.edu
http://www4.uwsp.edu/biology/faculty/RSchmitz/

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I'd like to hear from people who have or use a table-top or other
small compact SEM on a regular/routine basis. I'm interested to hear
how much you like it and how much use it gets...pros/cons, all the
dirt and all the gravy.
thanks,
Beth

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Hi,

When the beam is tilted it should stay on the same point on the sample
and only its angle change.
When the beam is shifted it should keep the same angle and only its
position change.
To assume that both conditions two stages of deflectors are needed. When
upper coils deflecte the beam to tilt it, lower coils deflecte the beam
to compensate the shift induced by the upper coils. When lower coils
deflecte the beam to shift it, upper coils deflecte the beam to
compensate the tilt induced by the lower coils. In case of modern JEOL
TEM ratios of compensation are adjustable with wobblers.
The link : http://www.matter.org.uk/tem/scanning_tilting_alignment.htm
gave by Marco is a good illustration concerning shifting but it is
faulty concerning tilting (lower coils are inactive).
Best greetings

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

-------- Message original --------
Sujet: [Microscopy] Re: viaWWW:difference between "Shift" and "Bright
Tilt"

Hello,
to centre the beam on the centre of the screen, just use the SHIFT wheel. It will translate=shift the beam.

If you need to adjust the tilt of the voltage or current axis of the beam to run in parallel to the optical axis, then activate with the "Bright Tilt" button the multifunctional wheel "Def/Stig", and properly adjust=tilt the axis with this. (see manual chapter 5.3.5 for further instructions on this).

To understand what�s happing, please have a look on i.e. this:
http://www.matter.org.uk/tem/scanning_tilting_alignment.htm

You will see that shifting does not only alter the CLA2, but also affects CLA1. They both (CLA1 and CLA2) are acting concerted, in certain relation to each other, on order obtain the translation of the beam.

On your JEM-2100F, the CLA1 effect might appear so small, that you can�t read it out easily from the Lens/Def Voltage Value panel. But if you write down the CLA values with the "Def/Lens Memory" tool before and after manipulating them, then you will see the effect well represented in the changing values.

Best greetings from the EM-Labs of CIC biomaGUNE,
Marco M�ller

----------------------------------------------------------------
Marco M�ller
Platform Manager - Electron Microscopy

CIC biomaGUNE
Parque Tecnol�gico de San Sebasti�n
Paseo Miram�n, 182. Ed. Empresarial C
20009 San Sebasti�n (Guip�zcoa)
SPAIN

Tel: +34 943 00 53 25
mmoller-at-cicbiomagune.es

-

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I have had good results with a Barnstead International "Super-Nuova" hot
plate. It comes in 7x7 and 10x10 inch sizes, also 100, 120 or 220 volt
models.
Precise temp control and holds set temperature well.

Geoff

On 4/23/2012 5:33 PM, rschmitz-at-uwsp.edu wrote:
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} My Corning stir hotplates that I use for drying thick sections onto glass slides have seen better days.
} And there is next to no control much less any temperature measurement.
} We just turn them on, barely, then let the heat up. We test the heat by touching the hot plate.
} If you can count to 5 before you have to remove your finger, then the plate is hot but not too hot.
} Does anyone have a hotplate that they would recommend I consider?
} Thanks for you help in advance
} Bob
} Dr. Robert J. Schmitz
} Associate Professor of Anatomical Sciences
} Department of Biology
} University of Wisconsin-Stevens Point
} 800 Reserve Street, 380 TNR Bld
} Stevens Point, WI 54481
} 715-346-2420
} Email: rschmitz-at-uwsp.edu
} http://www4.uwsp.edu/biology/faculty/RSchmitz/
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--
--
**********************************************
Geoff McAuliffe, Ph.D.
Neuroscience and Cell Biology
Robert Wood Johnson Medical School
675 Hoes Lane, Piscataway, NJ 08854
voice: (732)-235-4583
mcauliff-at-umdnj.edu
**********************************************

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Dear All, a very good day to you! My lab owns a JEOL 2010 TEM. Recently, the JEOL engineers have come to do a preventive�maintenance�for our system. They have change a new LaB6 filament and done a thorough check on any possible leakage and the power stability because before the�maintenance, our filament doesn't seems to be running in good condition. The filament current (we set at 104uA to 106 uA) drops�noticeably�faster to the background during usage and finally burn off (in a span of 4 months). While i am hoping that the�maintenance�would clean up the column and stop any possible leakage (they did detect a leak at V2, which is at around the camera chamber), the new filament which we are using this time doesn't look promising either.�

Firstly, before we allow the users to use (when tem is not in use), we observed that the filament is unstable �twice. Once is when the current suddenly drops from 101uA to zero when not in use. The second time is when the JEOL engineer is removing the sample holder (he is suppose to be an expert in his own machine). For the second case, the engineer told me that it is not because of any discharge from the filament by observing the change in the SIP.�Just from yesterday till today, the filament current drops from 106 uA (the engineer set at this current) back to 101uA �after a day of usage (many users came in to use this TEM). On the other hand, i try to observe the shape of the LaB6 filament under undersaturated condition. I did see 4 lines from different directions connecting to a dot (supposedly�it should be the tip, please correct me if i am wrong), but they seems to be tilted at an angle as the dot is not at the center of the beam observed under
the viewing screen. I try to use the gun tilt in attempt to bring the filament back to the center of the beam but it didn't seems to be working. What i observed is that the lines and the dots are fading fast into the beam (which also darkens) when i do the gun tilt.�

I would like to ask

1) If anyone has experience the fast dropping beam current during usage, if so what could be the possible reason
2) If i cannot bring the tip back to the center when using the gun tilt, what could have happen or what other alignment i could have use?

Pardon me because i am still learning how to use a TEM correctly.

Best Regards,
Yee Yan, Tay
FACTS Lab
Singapore

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7, 35 -- From: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
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Scanning Probe Microscopy for Energy Applications - September 12-13, Oak
Ridge, TN
*
*

*Scope *

Energy generation, storage, and conversion systems are an integral
component of emerging green technologies, including solar power,
automotive, and storage components of solar and wind energy economics.
Despite the rapidly expanding manufacturing capabilities and wealth of
phenomenological information on the macroscopic behaviors of energy
storage and conversion systems, the microscopic mechanisms underpinning
solar cell, battery and fuel cell operations in the nanometer to micron
range are not well understood. This series of keynote and invited talks
will cover the recent advances in characterization of energy relevant
materials systems using Scanning Probe Microscopy (SPM) techniques, as
well as the state of the art in energy dissipation and transformation
measurements by SPM. Topics include mapping of carrier dynamics and
photo-induced behavior of photovoltaic materials, ionic and electronic
transport in fuel cells and Li-ion batteries, energy harvesting, and
energy dissipation imaging by multiple resonant and band excitation
SPMs, as well material characterization using mass spectrometry combined
with SPM and optical spectroscopies for multimodal imaging. A number of
contributed talks will be included. Ultimately, our goal is to build a
network of materials scientists centered on the applications of SPM for
energy problems and to promote rapid dissemination of theoretical
knowledge, experimental protocols, and novel technique development in
this rapidly growing area.

The 3rd International Workshop on Scanning Probe Microscopy for Energy
Applications will be hosted in conjunction with the Annual Symposium of
the Tennessee Valley Chapter (TVC) of AVS. The TVC-AVS focus session is
an interdisciplinary meeting to discuss advances in SPM and applications
of nanostructure analysis. Contributions from all areas of nanoscience
and vacuum science are encouraged.

*Invited Speakers (partial list)*

·Richard Caprioli, Vanderbilt University – /Multi-modal imaging:
Combining MALDI-MS imaging with MRI and microscopy/

·Phillip First, Georgia Institute of Technology//– /Measuring the
effects of scalar and vector potentials in graphene/

·M. Hersam, Northwestern University – /Scanning probe microscopy of
energy materials/

·J. Huang, Sandia National Laboratory – /In-situ electrochemistry with
nanometer resolution/

·Lincoln Lauhon, Northwestern University – /Correlated functional
imaging of energy interconversion in hybrid Nanostructures /

·M. Pan, Oak Ridge National Laboratory – /Oxide surfaces on atomic
scale: STM quest/

·V. Sethuraman, Brown University – /Strain dynamics in energy materials/

·Z.L. Wang, Georgia Institute of Technology – /Nanopiezotronics and
nanowire based energy harvesting/

·Nick Winograd, Pennsylvania State University – /Nanoscale chemical
imaging of biomaterials with mass spectrometry/

·Vasilia Zorba, Lawrence Berkeley National Laboratory – /Optical near
and far-field femtosecond laser ablation for nanoscale chemical imaging/

*Call for Abstracts—Talks and Posters*

Both oral and poster presentations are welcome. A limited number of
graduate student fellowships will be awarded to reimburse the
registration fee and lodging at the conference hotel. Abstracts must be
received by August 13. Instructions and on-line submission will be
provided at the CNMS web site, www.cnms.ornl.gov
{http://www.cnms.ornl.gov} . Contact An-Ping Li (apli-at-ornl.gov
{mailto:apli-at-ornl.gov} ) for additional information on abstract
submission, Olga Ovchinnikova (ovchinnikovo-at-ornl.gov
{mailto:ovchinnikovo-at-ornl.gov} ) for student fellowship.

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18, 19 -- From: "Sergei V. Kalinin" {sergei2-at-ornl.gov}
18, 19 -- User-Agent: Mozilla/5.0 (Windows NT 6.1; WOW64; rv:11.0) Gecko/20120327 Thunderbird/11.0.1
18, 19 -- MIME-Version: 1.0
18, 19 -- To: microscopy-at-microscopy.com, sv9-at-ornl.gov
18, 19 -- Subject: Scanning Probe Microscopy for Energy Applications - September 12-13,
18, 19 -- Oak Ridge, TN
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Dear All: Our microscopy lab has some job openings for degree holders. The job�descriptions�are as below

Job description
1) � �TEMs�(Transmission Electron Microscopes)
a. � �Training�of new TEM user
b. � �Training�of TEM sample preparation equipment (includes Ultrasonic Disc Cutter, Disc
Grinder, Dimpler, Precision Ion Polishing System, Evaporator and Microtome)
c. � �Maintenance, routine check, alignment and calibration of TEM

2) � �Lab�routine running and management
a. � �Routine�lab work including overall taking care of lab equipment, service and troubleshooting
b. � �Managing�lab expenditure, consumables, accessories and equipment purchasing
c. � �Supporting to set up the equipment usage regulation/application

You can email me : YYTAY-at-ntu.edu.sg if you are interested in applying and i can forward to the person in charge.

Thank you!

Regards,
Yee Yan, Tay
FACTS Lab (http://facts.ntu.edu.sg)
Nanyang Technological University, Singapore

==============================Original Headers==============================
7, 35 -- From one_twinklestar-at-yahoo.com.sg Tue Apr 24 20:32:46 2012
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7, 35 -- Date: Wed, 25 Apr 2012 09:32:43 +0800 (SGT)
7, 35 -- From: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
7, 35 -- Reply-To: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
7, 35 -- Subject: Job Posting -at- Electron Microscopy Lab
7, 35 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
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-------- Original Message --------

Ralph Albrecht at Wisconsin was (is?) working on colloidal probes
with different morphologies (triangles, faceted, etc) to provide
distinguishable BSE signals. Some of the probes were pretty big, but a
number of them were { 15-18nm.

Ref: Meyer DA, Oliver JA, Albrecht RM (2010). Colloidal palladium
particles of different shapes for electron microscopy labeling. Microsc
Microanal. 16(1):33-42. Pubmed link:
http://www.ncbi.nlm.nih.gov/pubmed/20030909 .

Aaron
----------------------------
Aaron Barnes
Dunny Lab | Dept of Microbiology | U of Minnesota

} } Reply-To: Lee Cohen-Gould {lcgould-at-med.cornell.edu}
} } Below is the result of your form, submitted on Friday, April 20,
} } 2012 at 09:10:50 AM.
} }
} } realname - Lee Cohen-Gould
} } Email - lcgould-at-med.cornell.edu
} } EDUCATION - Graduate College
} } LOCATION - USA
} } SUBJECT_OF_QUESTION - multiple labels in immunogold
} } QUESTION - Hi All-
} } Nestor's filters keep rejecting me, so I'm coming in the back door.
} } I have a client who hopes to do triple immunogold labeling. I have
} } been noodling around with ideas including peroxidase-DAB or
} } peroxidase-ENZMET for one of the labels and 2 sizes of gold for the
} } others. I thought that I'd toss this out as an intellectual
} } exercise to those of you who do this all the time. Here's another
} } complication: she has 3 primary antibodies and all are available
} } from rabbit or goat hosts, but that's it. Two of the primaries will
} } have to be from the same host. I thought that I'd try Protein A or
} } G for one of them, using excess, untagged protein to block before
} } the second primary is used. Long and tedious. Any better ideas?
} } I know I can count on THE LIST for suggestions.
} } Thanks in advance!
} } Lee
} Login Host: 98.26.94.6
} Listserver Email Form V - 20120416
} ---------------------------------------------------------------------------
}
}
} ==============================Original Headers==============================
} 21, 28 -- From microscopylistserver-noreply-at-microscopy.com Tue Apr 24 21:01:50 2012
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==============================Original Headers==============================
4, 21 -- From barnesa-at-umn.edu Tue Apr 24 21:24:06 2012
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4, 21 -- Date: Tue, 24 Apr 2012 21:24:01 -0500
4, 21 -- From: Aaron Barnes {barnesa-at-umn.edu}
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Dear All, my apology, it is actually one job opening with the following job descriptions

1) � �TEMs�(Transmission Electron Microscopes)�
-Training�of new TEM user�
-Training�of TEM sample preparation equipment (includes Ultrasonic Disc Cutter, Disc
� � � � �Grinder, Dimpler, Precision Ion Polishing System, Evaporator and Microtome)�
-Maintenance, routine check, alignment and calibration of TEM

2) � �Lab�routine running and management�
-�Routine�lab work including overall taking care of lab equipment, service and troubleshooting
-�Managing�lab expenditure, consumables, accessories and equipment purchasing
-�Supporting to set up the equipment usage regulation/application�

My apology if there is any misunderstanding.

Cheers,
Yee Yan, Tay
FACTS Lab
NTU

----- Original Message -----
X-from: "one_twinklestar-at-yahoo.com.sg" {one_twinklestar-at-yahoo.com.sg}
To: one_twinklestar-at-yahoo.com.sg
Cc:
Sent: Wednesday, 25 April 2012, 9:33

Dear All: Our microscopy lab has some job openings for degree holders. The job�descriptions�are as below

Job description
1) � �TEMs�(Transmission Electron Microscopes)
a. � �Training�of new TEM user
b. � �Training�of TEM sample preparation equipment (includes Ultrasonic Disc Cutter, Disc
Grinder, Dimpler, Precision Ion Polishing System, Evaporator and Microtome)
c. � �Maintenance, routine check, alignment and calibration of TEM

2) � �Lab�routine running and management
a. � �Routine�lab work including overall taking care of lab equipment, service and troubleshooting
b. � �Managing�lab expenditure, consumables, accessories and equipment purchasing
c. � �Supporting to set up the equipment usage regulation/application

You can email me : YYTAY-at-ntu.edu.sg if you are interested in applying and i can forward to the person in charge.

Thank you!

Regards,
Yee Yan, Tay
FACTS Lab (http://facts.ntu.edu.sg)
Nanyang Technological University, Singapore

==============================Original Headers==============================
7, 35 -- From one_twinklestar-at-yahoo.com.sg Tue Apr 24 20:32:46 2012
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7, 35 -- From: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
7, 35 -- Reply-To: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
7, 35 -- Subject: Job Posting -at- Electron Microscopy Lab
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"That now leaves the problem of three targets with two hosts for the
primary antibodies. You could
use a biotinylated secondary antibody and labeled streptavidin as a
tertiary probe for one (suggest
the first one, detected with HRP-DAB), and block afterwards with excess
unlabeled secondary if
necessary. Then the other two targets can be treated as a double labeling
experiment, making sure
that the labeled secondaries are pre-absorbed against the host animals for
the other antibodies."

Hi, Rick:

In the approach your described (copied above), I assume the purpose of
blocking with unlabeled secondary after the first labeling using
immunoperoxidase (ABC system) is to block the epitopes on the first
primary (e.g. rabbit primary I) that are not bound by the first
anti-rabbit secondary (biotinylated). If that is the case, I think the
followings are potential issues.

1. Since the primary against one of the remaining targets (e.g. rabbit
primary II) is from the same species as the first primary (rabbit primary
I), rabbit primary II will bind to free Fab from the first secondary as
well as to the unlabeled secondary used for blocking.

2. Since two anti-rabbit secondaries are from different sources, they
probably are against different set of epitopes on rabbit primary. This
means that the second anti-rabbit secondary could still bind to the first
primary even with the blocking using unlabeled secondary.

I remember reading literature years ago that the reaction product of the
HRP could mask the antibody binding sites, although I have never tried it
myself. If that is the case, the HRP enzyme reaction may render the first
anti-rabbit secondary unavailable, which is helpful in reducing
cross-binding. Similarly, silver enhancement of ultrasmall gold
conjugates also have the effect of encapsulating the antibody therefore
double labeling using primaries of the same species can be done
sequentially (Electron microscopic immunocytochemistry. Silver enhancement
of colloidal gold marker allows double labeling with the same primary
antibody. Bienz K J Histochem Cytochem. 1986 Oct;34(10):1337-42.)

Thank you.

Hong

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"That now leaves the problem of three targets with two hosts for the
primary antibodies. You could use a biotinylated secondary antibody and
labeled streptavidin as a tertiary probe for one (suggest the first one,
detected with HRP-DAB), and block afterwards with excess unlabeled
secondary if necessary. Then the other two targets can be treated as a
double labeling experiment, making sure that the labeled secondaries are
pre-absorbed against the host animals for the other antibodies."

Hi, Rick:

In the approach your described (copied above), I assume the purpose of
blocking with unlabeled secondary after the first labeling using
immunoperoxidase (ABC system) is to block the epitopes on the first
primary (e.g. rabbit primary I) that are not bound by the first
anti-rabbit secondary (biotinylated). If that is the case, I think the
followings are potential issues.

1. Since the primary against one of the remaining targets (e.g. rabbit
primary II) is from the same species as the first primary (rabbit primary
I), rabbit primary II will bind to free Fab from the first secondary as
well as to the unlabeled secondary used for blocking.

2. Since two anti-rabbit secondaries are from different sources, they
probably are against different set of epitopes on rabbit primary. This
means that the second anti-rabbit secondary could still bind to the first
primary even with the blocking using unlabeled secondary.

I remember reading literature years ago that the reaction product of the
HRP could mask the antibody binding sites, although I have never tried it
myself. If that is the case, the HRP enzyme reaction may render the first
anti-rabbit secondary unavailable, which is helpful in reducing
cross-binding. Similarly, silver enhancement of ultrasmall gold
conjugates also have the effect of encapsulating the antibody therefore
double labeling using primaries of the same species can be done
sequentially (Electron microscopic immunocytochemistry. Silver enhancement
of colloidal gold marker allows double labeling with the same primary
antibody. Bienz K J Histochem Cytochem. 1986 Oct;34(10):1337-42.)

Thank you.

Hong

P.S. I am resending this reply because there was a formatting issue in
last one. Thank you again.

On 4/24/12 10:03 PM, "microscopylistserver-noreply-at-microscopy.com"
{microscopylistserver-noreply-at-microscopy.com} wrote:

}
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Dear Listers,
�
I need to label a membrane-bound protein (CAM or cell-adhesion molecule). I already tried
- 2 different fixations: 4% PFA for 15 min followed by 0,1% Triton x-100 10 min for permeabilization OR Methanol/acetone for 10 min at -20�C
- 3 different primaries (1 mono and 2 polyclonals)
- Blocking for 30 min with 1%BSA/PBS looks good
- Incubation with Ab for 1h at 37�C.
�
I seem to get a very faint signal with one primary and methanol/acetone fixation but I am near the limit of detection.
I wondered if someone had experience with IF on membrane-bound proteins, I have plenty of experience with IF on nuclear proteins but not these one.
Is there a special procedure I can follow to reveal an epitope embedded in the cell membrane?
I will try to increase the primary concentration and incubation time but in the meantime I am looking for other strategies to increase the signal.
I would prefer to go on with a methanol/acetone fixation since this strategy was the only one to give me a (low) signal.
�
Thank you for your suggestions
�
Stephane

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Dear listers,

One researcher came to our lab and asked a favor if I could view her research samples so that she could see the bacteria (Sporosarcina Pasteurii) and take a micrograph of it. For the samples that involves cementation + bacteria, I used a peltier stage (ESEM mode, no preparation) and I was able to get a favorable result. However, for the sample which supposedly contains thousands of bacteria (fresh / without cementation), I was not able to see even one!

Any idea how I could see these bacteria? Do we need to stain them? If yes, any staining recipe you could share? Sorry for the volume of questions. This is the first time I had biological samples to work at.

Thanks in advance.

Regards,
Melina Miralles

==============================Original Headers==============================
10, 33 -- From mmiralles-at-pi.ac.ae Wed Apr 25 04:39:56 2012
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10, 33 -- Subject: Viewing Bacteria
10, 33 -- Thread-Topic: Viewing Bacteria
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Melina,

What did you see? A more or less amorphous lump of goo or the like?
If so, you may have been seeing the biofilm, which will hide the
bacteria, even if thin. A higher accelerating voltage might indicate
bacteria below the film, but they wouldn't be clearly imaged.
The bacteria would need washing, or fix and dehydrate to remove the
biofilm water, in order to see the bacteria. Staining should not be
necessary. Although an OsO4 vapor fix might react with the bacteria
and not the sugars of the biofilm, which would help make the bacteria
visible, particularly in composition-mode backscatter imaging.

Phil

} Dear listers,
}
} One researcher came to our lab and asked a favor if I could view her
} research samples so that she could see the bacteria (Sporosarcina
} Pasteurii) and take a micrograph of it. For the samples that
} involves cementation + bacteria, I used a peltier stage (ESEM mode,
} no preparation) and I was able to get a favorable result. However,
} for the sample which supposedly contains thousands of bacteria
} (fresh / without cementation), I was not able to see even one!
}
} Any idea how I could see these bacteria? Do we need to stain them?
} If yes, any staining recipe you could share? Sorry for the volume of
} questions. This is the first time I had biological samples to work
} at.
}
} Thanks in advance.
}
} Regards,
} Melina Miralles

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

==============================Original Headers==============================
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Hi,
Does anyone know of techniques to locate pancreatic islets in epoxy
resin sections? They are sparse so I will try any stain that might help.
I have just used my standard Toluidine Blue and examine with a 40X
objective.

Thanks,
Larry

--
Larry Ackerman, Specialist
Electron Microscopy Lab
Manager, Diabetes& Endocrinology Research Center Microscopy Core
UCSF, Dept. of Anatomy, Rm S657
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu

415-999-4758

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Dear listers,

One researcher came to our lab and asked a favor if I could view her research samples so that she could see the bacteria (Sporosarcina Pasteurii) and take a micrograph of it. For the samples that involves cementation + bacteria, I used a peltier stage (ESEM mode, no preparation) and I was able to get a favorable result. However, for the sample which supposedly contains thousands of bacteria (fresh / without cementation), I was not able to see even one!

Any idea how I could see these bacteria? Do we need to stain them? If yes, any staining recipe you could share? Sorry for the volume of questions. This is the first time I had biological samples to work at.

Thanks in advance.

Regards,
Melina Miralles

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Hi Ashley,

You can always try a negative stain sample prep. It's an extremely fast way to view bacteria, viruses, ect. without having to chemically fix and process anything. You will need the negative stain, usually a 2+ACU- neutralized PTA (phototungstic acid), and some formvar carbon coated grids. What is the source of your bacteria, is it in suspension?

Scott Fish
EM Lab
California Animal Health and Food Safety Lab
1233 Thurman Lab
School of Vet. Med.
University of California, Davis
Davis, CA 95616
530-752-8760

-----Original Message-----
X-from: microscopylistserver-noreply+AEA-microscopy.com +AFs-mailto:microscopylistserver-noreply+AEA-microscopy.com+AF0-
Sent: Wednesday, April 25, 2012 6:20 AM
To: Fish, Scott S.

Dear Lee,

Triple labeling experiments can be done of course, but as was illustrated by Hong and commented by mr Powell, there are risks and pitfalls, especially when working with secondary antibodies. It is certainly not for the fainthearted and one would have to be lucky to get it right without some (and potentially a lot of) fiddling. The amount of controls involved can be mind-boggling. Blocking with a relatively high concentration of secondary antibody, tagged or not, brings its own risk of increased background and leaves a chance of cross-labeling, I do not think it can ever be absolute.

If going with secondary antibodies I would choose a different approach: perform two or three double labeling experiments instead as follows:

* antigen 1 and 2
* antigen 2 and 3
* antigen 1 and 3

Such an approach would in my view be imperative as a preparative step for validation of a triple labeling. The suggestions that were given for preventing cross labeling of antigens by blocking out epitopes on primary and/or secondary (DAB polymer, silver or gold enhancement) are more easily tested in a two-by-two set up. Mutatis mutandis, first of all the single labeling has to be established to work well for each individual antigen.

The protein A approach for multiple labeling worked out in the 80's and 90's by Hans Geuze's group in Utrecht is a neat way to overcome the issues caused by a secondary antibody approach when having primaries of the same species. An intermediate step of free protein A (200 �g/ml) after the smaller particle size protein A gold conjugate incubation will efficiently block the first primaries. Free binding sites on either protein A-gold or the blocking free protein A will of course bind the second primary from the same species, but since an antibody has only one binding site for a protein A molecule, this is not a problem. This should work in triple labeling, even with three primaries from one species and if my memory doesn't fail me, the Geuze group has demonstrated this as well.

Sorry Lee, it IS long and tedious no matter how you approach this :-)

Good luck from a magnificent autumn in New Zealand,

Jan

===

Jan Leunissen
Aurion ImmunoGold Reagents

e: leunissen-at-aurion.nl
i: www.aurion.nl

On 21/04/2012, at 4:14 AM, oshel1pe-at-cmich.edu wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} } Reply-To: Lee Cohen-Gould {lcgould-at-med.cornell.edu}
} } Below is the result of your form, submitted on Friday, April 20,
} } 2012 at 09:10:50 AM.
} }
} } realname - Lee Cohen-Gould
} } Email - lcgould-at-med.cornell.edu
} } EDUCATION - Graduate College
} } LOCATION - USA
} } SUBJECT_OF_QUESTION - multiple labels in immunogold
} } QUESTION - Hi All-
} } Nestor's filters keep rejecting me, so I'm coming in the back door.
} } I have a client who hopes to do triple immunogold labeling. I have
} } been noodling around with ideas including peroxidase-DAB or
} } peroxidase-ENZMET for one of the labels and 2 sizes of gold for the
} } others. I thought that I'd toss this out as an intellectual
} } exercise to those of you who do this all the time. Here's another
} } complication: she has 3 primary antibodies and all are available
} } from rabbit or goat hosts, but that's it. Two of the primaries will
} } have to be from the same host. I thought that I'd try Protein A or
} } G for one of them, using excess, untagged protein to block before
} } the second primary is used. Long and tedious. Any better ideas?
} } I know I can count on THE LIST for suggestions.
} } Thanks in advance!
} } Lee

==============================Original Headers==============================
21, 21 -- From leunissen-at-aurion.nl Thu Apr 26 00:20:52 2012
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I agree with Jan,
�
I tried the approach detailed by Rick some 15 years ago for immunofluorescence and it didn't give satisfactory results. It is very tricky to go around all cross-reactions.
As Jan points out, making all the controls necessary may be very tedious and we shouldn't forget that we are taking immuno-gold in TEM here, which is much more time-consuming than IF.
I suggested direct labeling of the primaries but it has its drawbacks too.
�
Regards,
Stephane
�
PS: If you want to visit Vienna and don't want to mess with mass-tourism of the summer monthes, do it the coming week-end. We'll get summer temperatures and a blue blue sky!
�
�
X-from: "leunissen-at-aurion.nl" {leunissen-at-aurion.nl}
To: nizets2-at-yahoo.com
Sent: Thursday, April 26, 2012 7:26 AM

Dear Lee,

Triple labeling experiments can be done of course, but as was illustrated by Hong and commented by mr Powell, there are risks and pitfalls, especially when working with secondary antibodies. It is certainly not for the fainthearted and one would have to be lucky to get it right without some (and potentially a lot of) fiddling. The amount of controls involved can be mind-boggling. Blocking with a relatively high concentration of secondary antibody, tagged or not, brings its own risk of increased background and leaves a chance of cross-labeling, I do not think it can ever be absolute.

If going with secondary antibodies I would choose a different approach: perform two or three double labeling experiments instead as follows:

* antigen 1 and 2
* antigen 2 and 3
* antigen 1 and 3

Such an approach would in my view be imperative as a preparative step for validation of a triple labeling. The suggestions that were given for preventing cross labeling of antigens by blocking out epitopes on primary and/or secondary (DAB polymer, silver or gold enhancement) are more easily tested in a two-by-two set up. Mutatis mutandis, first of all the single labeling has to be established to work well for each individual antigen.

The protein A approach for multiple labeling worked out in the 80's and 90's by Hans Geuze's group in Utrecht is a neat way to overcome the issues caused by a secondary antibody approach when having primaries of the same species. An intermediate step of free protein A (200 �g/ml) after the smaller particle size protein A gold conjugate incubation will efficiently block the first primaries. Free binding sites on either protein A-gold or the blocking free protein A will of course bind the second primary from the same species, but since an antibody has only one binding site for a protein A molecule, this is not a problem. This should work in triple labeling, even with three primaries from one species and if my memory doesn't fail me, the Geuze group has demonstrated this as well.

Sorry Lee, it IS long and tedious no matter how you approach this :-)

Good luck from a magnificent autumn in New Zealand,

Jan

===

Jan Leunissen
Aurion ImmunoGold Reagents

e: leunissen-at-aurion.nl
i: www.aurion.nl

On 21/04/2012, at 4:14 AM, oshel1pe-at-cmich.edu wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor:� The Microscopy Society of America
} To� Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
}
} } Reply-To: Lee Cohen-Gould {lcgould-at-med.cornell.edu}
} } Below is the result of your form, submitted on Friday, April 20,
} } 2012 at 09:10:50 AM.
} }
} } realname - Lee Cohen-Gould
} } Email - lcgould-at-med.cornell.edu
} } EDUCATION - Graduate College
} } LOCATION - USA
} } SUBJECT_OF_QUESTION - multiple labels in immunogold
} } QUESTION - Hi All-
} } Nestor's filters keep rejecting me, so I'm coming in the back door.
} } I have a client who hopes to do triple immunogold labeling.� I have
} } been noodling around with ideas including peroxidase-DAB or
} } peroxidase-ENZMET for one of the labels and 2 sizes of gold for the
} } others.� I thought that I'd toss this out as an intellectual
} } exercise to those of you who do this all the time.� Here's another
} } complication:� she has 3 primary antibodies and all are available
} } from rabbit or goat hosts, but that's it.� Two of the primaries will
} } have to be from the same host.� I thought that I'd try Protein A or
} } G for one of them,� using excess, untagged protein to block before
} } the second primary is used.� Long and tedious.� Any better ideas?
} } I know I can count on THE LIST for suggestions.
} } Thanks in advance!
} } Lee

==============================Original Headers==============================
21, 21 -- From leunissen-at-aurion.nl Thu Apr 26 00:20:52 2012
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21, 21 -- Subject: Re: [Microscopy] Multiple labels in immunogold
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21, 21 -- From: Jan Leunissen {leunissen-at-aurion.nl}
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Ashley

if you haven't got agar in your lab, I assume the bacteria have come from somewhere else because agar is the standard growth medium.

You don't necessarily have to encapsulate the bacteria in agar it is often just a convenient way of handling a pellet of small cells without them breaking up during fixation, dehydration and embedding. I have only used it if I needed to retain the structure of a microbial colony intact or because the cells were likely to dispersse. Most samples of bacteria can just be spun down in a microcentrifuge between each stage of fixation to retain a pellet.

Good luck.

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: microscopylistserver-noreply-at-microscopy.com [microscopylistserver-noreply-at-microscopy.com]
Sent: 25 April 2012 14:19
To: Malcolm Haswell

Hello Ashley,
Using molten agar for bacteria sample preparation is quite simple. General rule is to mix equal volumes of bacterial pellet and molten agar. After agar gelification, the agar-bacteria gel is cut into the small cubes (approx 1mm x 1mm x 1mm). The cubes are then processed as pieces of tissue. The advantage of this is that you do not need to centrifuge the bacteria sample between next processing steps.
In our lab, we use a low-melting agarose instead of agar.

Our common procedure is following:
A: Make a 4% low-melting agarose in washing buffer or ddH2O in a Eppendorf tube. It can be stored for some time in a refrigerator.

B: The procedure
1. fix the bacteria sample (in Eppendorf tube)
2. wash the sample well (at least three times)
3. solidify 4% agarose gel and temper it to 30 oC or 37 oC in water bath
4. gently spin down the washed bacteria sample
5. remove as much as possible of the supernatant
6. estimate the volume of the sediment
7. add an equal volume of 4% agarose and gently mix (after this step, you can gently spin down the mixture)
8. let solidify the mixture
9. cut off the bottom of Eppendorf tube containing the agar-bacteria gel
10. take the agar-bacteria gel out and cut it into the small cubes with a razor blade
11. postfix the cubes in OsO4
12. process further as common tissue bloc (wash, dehydrate, embed into resin, etc.)

My best regards Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

On Wednesday 25 of April 2012 15:13:43 you wrote:
}
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} Title-Subject: Bacteria TEM sample prep
}
} Message: Can anyone explain how to prepare bacterial cells for TEM observation. The literature I'm
} reading mentions adding molten agar to the fixative, however, I've never come across agar in our
} lab. Can anyone comment on that? Is it ok to go from the fixative to the wash to the fixative? If
} anyone can answer this question or give ANY advice, from prepping grids, to embedding, will all be
} Highly appreciated. Many thanks!
} Ashley Rodriguez
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7, 23 -- From benada-at-biomed.cas.cz Thu Apr 26 03:41:36 2012
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Hello,
I am sorry, I have done a mistake in the step 3. of the procedure. It should be:

3. liquify 4% agarose gel and temper it to 30 oC or 37 oC in water bath

Oldrich

On Thursday 26 of April 2012 10:41:30 Oldrich Benada wrote:
} Hello Ashley,
} Using molten agar for bacteria sample preparation is quite simple. General rule is to mix equal volumes of bacterial pellet and molten agar. After agar gelification, the agar-bacteria gel is cut into the small cubes (approx 1mm x 1mm x 1mm). The cubes are then processed as pieces of tissue. The advantage of this is that you do not need to centrifuge the bacteria sample between next processing steps.
} In our lab, we use a low-melting agarose instead of agar.
}
}
} Our common procedure is following:
} A: Make a 4% low-melting agarose in washing buffer or ddH2O in a Eppendorf tube. It can be stored for some time in a refrigerator.
}
} B: The procedure
} 1. fix the bacteria sample (in Eppendorf tube)
} 2. wash the sample well (at least three times)
} 3. solidify 4% agarose gel and temper it to 30 oC or 37 oC in water bath
} 4. gently spin down the washed bacteria sample
} 5. remove as much as possible of the supernatant
} 6. estimate the volume of the sediment
} 7. add an equal volume of 4% agarose and gently mix (after this step, you can gently spin down the mixture)
} 8. let solidify the mixture
} 9. cut off the bottom of Eppendorf tube containing the agar-bacteria gel
} 10. take the agar-bacteria gel out and cut it into the small cubes with a razor blade
} 11. postfix the cubes in OsO4
} 12. process further as common tissue bloc (wash, dehydrate, embed into resin, etc.)
}
} My best regards Oldrich
}
} }
} } ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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} } Email: arodriguez334-at-students.deltacollege.edu
} } Name: Ashley Rodriguez
} }
} } Organization: SJDC EM
} }
} } Title-Subject: Bacteria TEM sample prep
} }
} } Message: Can anyone explain how to prepare bacterial cells for TEM observation. The literature I'm
} } reading mentions adding molten agar to the fixative, however, I've never come across agar in our
} } lab. Can anyone comment on that? Is it ok to go from the fixative to the wash to the fixative? If
} } anyone can answer this question or give ANY advice, from prepping grids, to embedding, will all be
} } Highly appreciated. Many thanks!
} } Ashley Rodriguez
} } San Joaquin Delta College EM Lab
} }
} } Login Host: 67.181.108.133
} } Listserver Email Form V - 20120416
} } ---------------------------------------------------------------------------
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4, 23 -- From benada-at-biomed.cas.cz Thu Apr 26 03:51:01 2012
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Linden Gledhill wants to grow snowflakes at home. I don't know if he has succeeded, but Wired

http://www.wired.com/wiredscience/2012/04/snowflake-growing-machine/

has some great images including a time lapsed photomicrograph video of what looks like hoarfrost growing. It's very cool (pun intended).

One of the things I find interesting is how scientist express themselves in their hobbies. (Who knew a cattle prod would be so useful?)

Stay safe.............
Frank

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Hi

From our experience, the answer is yes! Current plastic boxes or baggs
will outgaz solvents remaining from their manufacturing. I've no
quantitative data, only the conclusions of studies made by a collegue
here, now retird, made 20 years ago. His conclusion was that, if the
surface cleanless of a sample is important, it must be stored only in
clean (plasma treated, at that time by glow discharge) glassware. Never
plastic.
I observed too such contaminations, but cannot share more than
qualitative observations.

On the other hand, the semiconducteur industry uses most plastic boxes
(fluoroware). They cost a lot...
Maybe someone from that domain has more informations ?

Hope it helps

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Mat�riaux de Strasbourg
D�partement Surfaces et Interfaces
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail : Jacques.Faerber-at-ipcms.unistra.fr

Le 26/04/2012 03:11, microscopylistserver-noreply-at-microscopy.com a �crit :
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} Title-Subject: Specimen preparation - Carbon contamination in plastic bag
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} Can a semiconductor or dielectric material be contaminated by carbon by storing it in a plastic bag?
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Hello,

I'm looking for PCI interface cards for two SPOT Insight color
cameras, both model #3.2.0. I know Diagnostic Instruments sells them
on their website but I'm looking for used pricing.

Thanks,
Esteban

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Hi Beth

We also were seeking this information before making our decision last year. You might like to search the list but if I remember correctly the few valuable responses came privately. To put this in perspective, our application is diagnostic/biological.

In our experience it came down to budget, service (in Australia) and whether you want a toy or a tool (to put it bluntly in guttural terms). The table-top SEMs are indeed a very nice, easy to operate and meet a niche role but they didn't offer the control and magnification we were seeking. We ended up going for a small SEM. In short, it is important to know what you expect from the instrument to know if the table-top will meet that.

Interestingly, one supplier we looked at is pushing into the gap between toy and tool with one of its models and its offering is quite useful and described as "cute".

Happy to discuss our experience further off-list,

Regards
Naomi

} } } {beth-at-plantbio.uga.edu} 4/24/2012 8:40 am } } }

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I'd like to hear from people who have or use a table-top or other
small compact SEM on a regular/routine basis. I'm interested to hear
how much you like it and how much use it gets...pros/cons, all the
dirt and all the gravy.
thanks,
Beth

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Late reply...

I would like to know the trick. I have never been fully successful in
removing dried, crystallized colloidal silica from a polished surface.
What success I have experienced is by using a rich solution of Micro90
(no vested interest)/hot water in a high specific energy ultrasonic
cleaner. Usually need to lightly swab the surface too although this is
likely to damage the polish on many softer materials. ...Sometimes I
can "read between the lines" (scratches).

Beware that Micro90 (http://www.ipcol.com/pdfs/M90_msds.pdf) is strongly
alkaline and may etch some metals.

Of course the ideal solution, as you know, is to not let the silica bond
in the first place . To facilitate this, I never let the surface dry
before cleaning. Immediately upon leaving the polishing wheel, I apply a
layer of water/Micro90 then go to the previously mentioned ultrasonic
sequence. Don't swab, but multiple UT (M-90, then DI water) and finish
rinse with DI water then alcohol.

If none of this works, try a chisel ;)

Woody

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Hello Listers!

I want to let everyone know about a couple of opportunities in the Dallas area. I currently have an opening for a technologist position in our diagnostic EM lab. If you have diagnostic EM experience, I would highly encourage you to apply.
Below is the official line on the position:
We are looking for a dedicated E/M technologist to perform all aspects of electron microscopy, (processing, sectioning, staining slides, grids, and loading EM scope). Some experience in Immunocytochemistry is helpful. Digital imaging applications and light microscopy highly desired. Experience with histology, graphics, digital imaging and statistics a plus! Biological TEM certification from the Microscopy Society of America is preferred, but not necessary.

Interested candidates can learn more by visiting: http://www.medfusionservices.com/laboratoryjobs.html

Steve Lee, BS, HT(ASCP)
Manager of Electron Microscopy/Imaging
972-966-7176
slee-at-medfusionsvs.com

The Convergence Center
2501 S. STATE HWY 121, Suite 1100
Lewisville, TX 75067
www.medfusionservices.com

med fusion PRIVACY AND CONFIDENTIALITY NOTICE: The information transmitted in this message and any attachments is intended for the person or entity to which it is addressed and may contain proprietary, confidential, and/or privileged material, which may include individually identifiable health information that requires safeguarding in compliance with the Health Insurance Portability and Accountability Act of 1996 as well as other federal and state laws. Any review, duplication, retransmission, distribution, or other use of, or taking of any action in reliance upon this information by persons or entities other than the intended recipient is prohibited. If you have received this message in error, please notify the sender immediately by return e-mail and delete all of the information from all computers or systems.

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We are looking into using the Kleinschmidt technique of spreading monolayers of DNA molecules for TEM imaging.
Any hints and advices would be greatly appreciated. We do not have experience with this method and even basic info will be invaluable. It seams that the use of Langmuir trough is a necessary for successful monolayer spreading, it this correct?

Thank you,

Krassimir Bozhilov

Central Facility for Advanced Microscopy and Microanalysis - CNAS
Materials Science and Engineering Program - BCOE
University of California,
Riverside, CA 92521

phone 951 827 2998
fax 951 827 2489

==============================Original Headers==============================
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8, 33 -- Subject: DNS monolayer
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On Apr 27, 2012, at 4:54 AM, microscopylistserver-
noreply-at-microscopy.com wrote:

} Name: Michael Miller
}
} Organization: Auburn University
}
} Title-Subject: TEM negative scanner
}
} Message: For those of us still using 4489 film in their TEM, what is
} the latest and greatest
} negative scanner on the market today for handling this type of
} negative? I am in need of replacing
} my old Epson scanner Thanks for any help on this subject.

Dear Michael,
The answer will depend on the type of images you want to scan,
although for the usual images of conventional sections and for
qualitative results, the scanner you choose will likely be determined
by ease of use and reliability. If, however, you need quantitative
results and/or are scanning images with very high dynamic range, such
as diffraction patterns, you will want a scanner that gives the
optical density of each pixel independently of the values of the
surrounding pixels. In the former case, a flat-bed scanner is
adequate, but in the latter case, you would need a scanner that uses a
small spot of illumination and rasters that spot with respect to the
film. I am not familiar with what is currently available, so I will
let others give opinions on particular brands.
Yours,
Bill

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Krassimir

It's a few years since I had to do this but it did seem like quite an achievement because we got it to a stage where undergraduate classes brought their bacterial plasmid samples over to the lab and we got some pretty decent results without any major expenditure on the same day.

The main features of the technique are:
1. to get a high quality source of water - we used triple distilled and only stored in acid washed and thoroughly rinsed glass stoppered containers. Deionised water and any water stored with plastic may risk the presence of "surface active agents" which could severely interfere with the spreading technique.
2. A Langmuir Trough is recommended for the casting of films incorporating the DNA but we found various other dishes (eg glass petri dishes worked as well)
3. All glassware and reagents should be thoroughly acid washed and rinsed in triple distilled water BUT NOT DETERGENT (a surface active agent). I found it best to have these prepared and stored separately.
4. The spreading technique worked well enough for simple plasmids.
5. The shadowing technique is best done by rotary shadowing where the angle of shadowing is fairly critical for getting the best results. We didn't have a rotary shadowing unit so I simply shadowed the prepared films, rotated them by 90 degrees and shadowed again. We got some really nice results but in the end this became time consuming. We even tried to build our own rotary turntable but most motors are not designed to work in vacuum with metal spraying about.
6. Our results were checked at about 50k on the TEM but I tended to photograph at much lower magnifications. This was done because we could then enlarge by 5x or 10x on the final prints and cut the picture up into sections so every student got a unique sample of plasmids to measure.

Somewhere I have a complete schedule for the process. I wrote 2 versions:
one simpler version for the students which outlined the technique
a second version for me with all the preparation details.

This technique wasn't perfect but it did work for simple plasmid preparations and consistently produced results with little expenditure but you might find it useful to do a pilot study and consider expenditure on Langmuir Troughs and rotary shadowing units if necessary. If you want me to forward the full technique sheets and an example of the results please let me know.

Good luck

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk

________________________________________
X-from: krassimir.bozhilov-at-ucr.edu [krassimir.bozhilov-at-ucr.edu]
Sent: 27 April 2012 20:30
To: Malcolm Haswell

We are looking into using the Kleinschmidt technique of spreading monolayers of DNA molecules for TEM imaging.
Any hints and advices would be greatly appreciated. We do not have experience with this method and even basic info will be invaluable. It seams that the use of Langmuir trough is a necessary for successful monolayer spreading, it this correct?

Thank you,

Krassimir Bozhilov

Central Facility for Advanced Microscopy and Microanalysis - CNAS
Materials Science and Engineering Program - BCOE
University of California,
Riverside, CA 92521

phone 951 827 2998
fax 951 827 2489

==============================Original Headers==============================
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8, 33 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
8, 33 -- Subject: DNS monolayer
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19, 36 -- From malcolm.haswell-at-sunderland.ac.uk Mon Apr 30 06:44:20 2012
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19, 36 -- From: Malcolm Haswell {malcolm.haswell-at-sunderland.ac.uk}
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Dear All,

I just wanted to thank you for the answers and suggestions received
regarding our problem with distortions of SEM images. We will be
investigating the problem this summer (now that my teaching semester
ended!) following the many useful advices we received.

Sincerely

Luca Fedele

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
----------------------

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A.Word.A.Day
with Anu Garg

nestor

PRONUNCIATION:
(NES-tuhr)

MEANING:
noun: A wise old man.

ETYMOLOGY:
X-from Nestor, king of Pylos, who was the oldest and wisest of the Greeks and served as a counselor in the Trojan War. Earliest documented use: around 1510.

USAGE:
"Roland Shaw was not only an oil man; he was the Nestor of the oil business, there when the first donkey nodded."
Bruce Anderson; The Long-Life Cocktail; The Spectator (London, UK); Nov 19, 2011.

Thanks, Nestor, for being our wise guy!!

cheers

Dr. S. Shea Miller
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
960 Carling Avenue | 960, avenue Carling
Ottawa, ON K1A 0C6
E-mail Address / Adresse courriel shea.miller-at-agr.gc.ca
Telephone | T�l�phone 613-759-1760
Facsimile | T�l�copieur 613-759-1701
Teletypewriter | T�l�imprimeur 613-773-2600
Government of Canada | Gouvernement du Canada

==============================Original Headers==============================
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Epic!

Al Coritz
Applications & Service Manager
EMS Technical Service
www.emsdiasum.com

-----Original Message-----
X-from: Shea.Miller-at-AGR.GC.CA [mailto:Shea.Miller-at-AGR.GC.CA]
Sent: Tuesday, May 01, 2012 9:25 AM
To: Sampleprep-at-earthlink.net

A.Word.A.Day
with Anu Garg

nestor

PRONUNCIATION:
(NES-tuhr)

MEANING:
noun: A wise old man.

ETYMOLOGY:
X-from Nestor, king of Pylos, who was the oldest and wisest of the Greeks
and served as a counselor in the Trojan War. Earliest documented use: around
1510.

USAGE:
"Roland Shaw was not only an oil man; he was the Nestor of the oil business,
there when the first donkey nodded."
Bruce Anderson; The Long-Life Cocktail; The Spectator (London, UK); Nov 19,
2011.

Thanks, Nestor, for being our wise guy!!

cheers

Dr. S. Shea Miller
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
960 Carling Avenue | 960, avenue Carling Ottawa, ON K1A 0C6 E-mail Address /
Adresse courriel shea.miller-at-agr.gc.ca Telephone | T�l�phone 613-759-1760
Facsimile | T�l�copieur 613-759-1701 Teletypewriter | T�l�imprimeur
613-773-2600 Government of Canada | Gouvernement du Canada

==============================Original Headers==============================
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Excellent!

But... "When the first donkey nodded" ??? Can
our wise Nestor, or anyone, explain that one??

TB
}
} A.Word.A.Day
} with Anu Garg
}
} nestor
}
}
} PRONUNCIATION:
} (NES-tuhr)
}
} MEANING:
} noun: A wise old man.
}
} ETYMOLOGY:
} X-from Nestor, king of Pylos, who was the oldest
} and wisest of the Greeks and served as a
} counselor in the Trojan War. Earliest documented
} use: around 1510.
}
} USAGE:
} "Roland Shaw was not only an oil man; he was the
} Nestor of the oil business, there when the first
} donkey nodded."
} Bruce Anderson; The Long-Life Cocktail; The
} Spectator (London, UK); Nov 19, 2011.
}
} Thanks, Nestor, for being our wise guy!!
}
} cheers
}
}
} Dr. S. Shea Miller
} ECORC | CRECO
} Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
} 960 Carling Avenue | 960, avenue Carling
} Ottawa, ON K1A 0C6
} E-mail Address / Adresse courriel shea.miller-at-agr.gc.ca
} Telephone | T�l�phone 613-759-1760
} Facsimile | T�l�copieur 613-759-1701
} Teletypewriter | T�l�imprimeur 613-773-2600
} Government of Canada | Gouvernement du Canada
}
}

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \
University of Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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A "nodding donkey" refers to an oil pump because of the resemblance to a donkey's head going up and down as he/she each grass. They are also sometimes called "thirsty birds"

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: baskin-at-bio.umass.edu [mailto:baskin-at-bio.umass.edu]
Sent: Tuesday, May 01, 2012 10:00 AM
To: Phillips, Thomas E.

Excellent!

But... "When the first donkey nodded" ??? Can
our wise Nestor, or anyone, explain that one??

TB
}
} A.Word.A.Day
} with Anu Garg
}
} nestor
}
}
} PRONUNCIATION:
} (NES-tuhr)
}
} MEANING:
} noun: A wise old man.
}
} ETYMOLOGY:
} X-from Nestor, king of Pylos, who was the oldest and wisest of the
} Greeks and served as a counselor in the Trojan War. Earliest documented
} use: around 1510.
}
} USAGE:
} "Roland Shaw was not only an oil man; he was the Nestor of the oil
} business, there when the first donkey nodded."
} Bruce Anderson; The Long-Life Cocktail; The Spectator (London, UK); Nov
} 19, 2011.
}
} Thanks, Nestor, for being our wise guy!!
}
} cheers
}
}
} Dr. S. Shea Miller
} ECORC | CRECO
} Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire
} Canada
} 960 Carling Avenue | 960, avenue Carling Ottawa, ON K1A 0C6 E-mail
} Address / Adresse courriel shea.miller-at-agr.gc.ca Telephone | T�l�phone
} 613-759-1760 Facsimile | T�l�copieur 613-759-1701 Teletypewriter |
} T�l�imprimeur 613-773-2600 Government of Canada | Gouvernement du
} Canada
}
}

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \
University of Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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16, 31 -- From PhillipsT-at-missouri.edu Tue May 1 10:03:28 2012
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I think the nodding donkey refers to the small oil wells, which look remarkably like nodding donkeys dotting fields....

Dr. S. Shea Miller
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
960 Carling Avenue | 960, avenue Carling
Ottawa, ON K1A 0C6
E-mail Address / Adresse courriel shea.miller-at-agr.gc.ca
Telephone | T�l�phone 613-759-1760
Facsimile | T�l�copieur 613-759-1701
Teletypewriter | T�l�imprimeur 613-773-2600
Government of Canada | Gouvernement du Canada

-----Original Message-----
X-from: Tobias Baskin [mailto:baskin-at-bio.umass.edu]
Sent: May-01-12 11:00 AM
To: Miller, Shea
Cc: microscopy-at-microscopy.com

Yee Yan,
I am mainly familiar with Kimball LaB6 cathodes in SEMs, but a couple of
things jumped out at me in your message.

A Kimball tip with the 15� flat should never be run above about 60�A or it
will fail prematurely. Check with the manufacturer of your tip for more
details.

Second, although I've had very little to do with TEMs over the years, most
typically run at much lower emission currents than SEMs, in the range of
10-15�A, I believe. What does your user's manual recommend?

The emission current could be the whole answer to your short filament life.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: one_twinklestar-at-yahoo.com.sg [mailto:one_twinklestar-at-yahoo.com.sg]
Sent: Tuesday, April 24, 2012 12:43 PM
To: kenconverse-at-qualityimages.biz

Dear All, a very good day to you! My lab owns a JEOL 2010 TEM. Recently, the
JEOL engineers have come to do a preventive�maintenance�for our system. They
have change a new LaB6 filament and done a thorough check on any possible
leakage and the power stability because before the�maintenance, our filament
doesn't seems to be running in good condition. The filament current (we set
at 104uA to 106 uA) drops�noticeably�faster to the background during usage
and finally burn off (in a span of 4 months). While i am hoping that
the�maintenance�would clean up the column and stop any possible leakage
(they did detect a leak at V2, which is at around the camera chamber), the
new filament which we are using this time doesn't look promising either.�

Firstly, before we allow the users to use (when tem is not in use), we
observed that the filament is unstable �twice. Once is when the current
suddenly drops from 101uA to zero when not in use. The second time is when
the JEOL engineer is removing the sample holder (he is suppose to be an
expert in his own machine). For the second case, the engineer told me that
it is not because of any discharge from the filament by observing the change
in the SIP.�Just from yesterday till today, the filament current drops from
106 uA (the engineer set at this current) back to 101uA �after a day of
usage (many users came in to use this TEM). On the other hand, i try to
observe the shape of the LaB6 filament under undersaturated condition. I did
see 4 lines from different directions connecting to a dot (supposedly�it
should be the tip, please correct me if i am wrong), but they seems to be
tilted at an angle as the dot is not at the center of the beam observed
under
the viewing screen. I try to use the gun tilt in attempt to bring the
filament back to the center of the beam but it didn't seems to be working.
What i observed is that the lines and the dots are fading fast into the beam
(which also darkens) when i do the gun tilt.�

I would like to ask

1) If anyone has experience the fast dropping beam current during usage, if
so what could be the possible reason
2) If i cannot bring the tip back to the center when using the gun tilt,
what could have happen or what other alignment i could have use?

Pardon me because i am still learning how to use a TEM correctly.

Best Regards,
Yee Yan, Tay
FACTS Lab
Singapore

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7, 35 -- From: Tay Yee Yan {one_twinklestar-at-yahoo.com.sg}
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7, 35 -- Subject: Lab6 Filament Problems
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20, 29 -- From kenconverse-at-qualityimages.biz Tue May 1 10:19:01 2012
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20, 29 -- "MSA Listserver" {microscopy-at-microscopy.com}
20, 29 -- Subject: RE: [Microscopy] Lab6 Filament Problems
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

We use Kimball LaB6 cathodes in our FEI CM20 TEM. Let me add a couple
of points to those made by Ken

1. Filament current - we typically run our below 40 microamps. It has
been quite while since I used a JEOL TEM, but if I recall correctly,
there is typically a dark current, so Ye Yan's 106 microamp current
may include the dark current - please let us know the value of the
emission current - dark current.

2. LaB6 cathodes need to be saturated slowly. I wrote a
DigitalMicrograph script to saturate my cathode over about 10 minutes
and do the required sample tracking chores while it runs. I should
also note that one should check saturation again after 10-15 minutes
because it can creep up. It is too easy to inadvertently over-saturate
the cathode. The cathode should be turned down slowly to minimize
thermal shock. I really wish our CM20 had a manual gun isolation valve
so I could close the valve with the cathode saturated during sample
exchange...

3. The cathode lifetime is also a function of vacuum. How good is your
gun vacuum?

Hope this helps,
John Minter
Eastman Kodak

On Tue, May 1, 2012 at 11:19 AM, {kenconverse-at-qualityimages.biz} wrote:
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: �The Microscopy Society of America
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} ----------------------------------------------------------------------------
}
} Yee Yan,
} I am mainly familiar with Kimball LaB6 cathodes in SEMs, but a couple of
} things jumped out at me in your message.
}
} A Kimball tip with the 15� flat should never be run above about 60�A or it
} will fail prematurely. �Check with the manufacturer of your tip for more
} details.
}
} Second, although I've had very little to do with TEMs over the years, most
} typically run at much lower emission currents than SEMs, in the range of
} 10-15�A, I believe. �What does your user's manual recommend?
}
} The emission current could be the whole answer to your short filament life.
}
} Ken Converse
} owner
}
} QUALITY IMAGES
} Servicing Scanning Electron Microscopes
} Since 1981
} 474 So. Bridgton Rd.
} Bridgton, ME �04009
} 207-647-4348
} kenconverse-at-qualityimages.biz
} qualityimages.biz
}
}
} -----Original Message-----
} X-from: one_twinklestar-at-yahoo.com.sg [mailto:one_twinklestar-at-yahoo.com.sg]
} Sent: Tuesday, April 24, 2012 12:43 PM
} To: kenconverse-at-qualityimages.biz
} Subject: [Microscopy] Lab6 Filament Problems
}
}
}
}
} ----------------------------------------------------------------------------
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}
} Dear All, a very good day to you! My lab owns a JEOL 2010 TEM. Recently, the
} JEOL engineers have come to do a preventive�maintenance�for our system. They
} have change a new LaB6 filament and done a thorough check on any possible
} leakage and the power stability because before the�maintenance, our filament
} doesn't seems to be running in good condition. The filament current (we set
} at 104uA to 106 uA) drops�noticeably�faster to the background during usage
} and finally burn off (in a span of 4 months). While i am hoping that
} the�maintenance�would clean up the column and stop any possible leakage
} (they did detect a leak at V2, which is at around the camera chamber), the
} new filament which we are using this time doesn't look promising either.
}
} Firstly, before we allow the users to use (when tem is not in use), we
} observed that the filament is unstable �twice. Once is when the current
} suddenly drops from 101uA to zero when not in use. The second time is when
} the JEOL engineer is removing the sample holder (he is suppose to be an
} expert in his own machine). For the second case, the engineer told me that
} it is not because of any discharge from the filament by observing the change
} in the SIP.�Just from yesterday till today, the filament current drops from
} 106 uA (the engineer set at this current) back to 101uA �after a day of
} usage (many users came in to use this TEM). On the other hand, i try to
} observe the shape of the LaB6 filament under undersaturated condition. I did
} see 4 lines from different directions connecting to a dot (supposedly�it
} should be the tip, please correct me if i am wrong), but they seems to be
} tilted at an angle as the dot is not at the center of the beam observed
} under
} �the viewing screen. I try to use the gun tilt in attempt to bring the
} filament back to the center of the beam but it didn't seems to be working.
} What i observed is that the lines and the dots are fading fast into the beam
} (which also darkens) when i do the gun tilt.
}
} I would like to ask
}
} 1) If anyone has experience the fast dropping beam current during usage, if
} so what could be the possible reason
} 2) If i cannot bring the tip back to the center when using the gun tilt,
} what could have happen or what other alignment i could have use?
}
} Pardon me because i am still learning how to use a TEM correctly.
}
} Best Regards,
} Yee Yan, Tay
} FACTS Lab
} Singapore
}
}
} ==============================Original Headers==============================
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Hi all,

I want to purchase a UPS system for our SEMs. Would appreciate information
as to vendors. Specifications we need are:
- Rating = 8 kVA / 6.4 kW
- Typical backup time of core instrument (i.e. at ca. 27% load) = 60 min.
- Input voltage = 172 - 285 Vac
- Input frequency = 40 - 70 Hz
- Output voltage = 230 Vac
- Output frequency = 50 / 60 Hz.

Any suggestions of vendors or internet sites that may have appropriate
systems would be appreciated.

Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

==============================Original Headers==============================
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Hi Debby,

APC has a wide range of selection (No commercial interest). You can check their range on this link:

http://www.apc.com/products/category.cfm?id=13

Hope this helps.

Sathya Srinivasan

Research Associate

Former employee of Experimental Imaging Center

(looking out for a job in microscopy/ image analysis)

University of Calgary

Calgary, AB, Canada

} Date: Tue, 1 May 2012 19:23:15 -0500
} To: sathya_sr70-at-hotmail.com
} From: dsherman-at-purdue.edu
} Subject: [Microscopy] UPS system
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi all,
}
} I want to purchase a UPS system for our SEMs. Would appreciate information
} as to vendors. Specifications we need are:
} - Rating = 8 kVA / 6.4 kW
} - Typical backup time of core instrument (i.e. at ca. 27% load) = 60 min.
} - Input voltage = 172 - 285 Vac
} - Input frequency = 40 - 70 Hz
} - Output voltage = 230 Vac
} - Output frequency = 50 / 60 Hz.
}
} Any suggestions of vendors or internet sites that may have appropriate
} systems would be appreciated.
}
} Debby
}
} ---
} Debby Sherman, Interim Director Phone: 765-494-6666
} Life Science Microscopy Facility FAX: 765-494-5896
} Purdue University E-mail: dsherman-at-purdue.edu
} S-052 Whistler Building
} 170 S. University Street
} West Lafayette, IN 47907
} http://www.ag.purdue.edu/facilities/microscopy
}
}
}
}
} ==============================Original Headers==============================
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} 8, 28 -- To: "message to: MSA list" {microscopy-at-microscopy.com}
} 8, 28 -- Date: Tue, 1 May 2012 20:13:26 -0400
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Dear All,
is there an easy way to make a gold on carbon test specimen for SEM ?

Thanks,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

==============================Original Headers==============================
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I have infiltrated Arabidopsis tissue with LR White resin and flat embedded
between two glass slides. I need to cure these specimens at 4 degree C
under UV light (I have a set of 2x15watt UV lamp setting). What is the
proper distance of UV lamps from specimen? an for what duration? These
lamps are almost 3-4 years or might be older, do I need to change them?

--
Katayoun Mansouri
Postdoctoral Research Scholar
Haigler Lab
Dept. of Crop Science
North Carolina State University
Raleigh, NC, 27695 U.S.A

==============================Original Headers==============================
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Dear All, thank you very much for all your nice advices! Our background current is 101 micro Amp. I have checked with JEOL, 106 micro Amp is their recommended beam current to be used in our machine. I should check with them the cathode. But i think its the brand from Denka. I will let you all know the details of the of the Lab6 filament.

Cheers,
Yee Yan, Tay

----- Original Message -----
X-from: "jrminter-at-rochester.rr.com" {jrminter-at-rochester.rr.com}
To: one_twinklestar-at-yahoo.com.sg
Cc:
Sent: Wednesday, 2 May 2012, 1:29

We use Kimball LaB6 cathodes in our FEI CM20 TEM. Let me add a couple
of points to those made by Ken

1. Filament current - we typically run our below 40 microamps. It has
been quite while since I used a JEOL TEM, but if I recall correctly,
there is typically a dark current, so Ye Yan's 106 microamp current
may include the dark current - please let us know the value of the
emission current - dark current.

2. LaB6 cathodes need to be saturated slowly. I wrote a
DigitalMicrograph script to saturate my cathode over about 10 minutes
and do the required sample tracking chores while it runs. I should
also note that one should check saturation again after 10-15 minutes
because it can creep up. It is too easy to inadvertently over-saturate
the cathode. The cathode should be turned down slowly to minimize
thermal shock. I really wish our CM20 had a manual gun isolation valve
so I could close the valve with the cathode saturated during sample
exchange...

3. The cathode lifetime is also a function of vacuum. How good is your
gun vacuum?

Hope this helps,
John Minter
Eastman Kodak

On Tue, May 1, 2012 at 11:19 AM,� {kenconverse-at-qualityimages.biz} wrote:
}
}
}
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} ----------------------------------------------------------------------------
}
} Yee Yan,
} I am mainly familiar with Kimball LaB6 cathodes in SEMs, but a couple of
} things jumped out at me in your message.
}
} A Kimball tip with the 15� flat should never be run above about 60�A or it
} will fail prematurely. �Check with the manufacturer of your tip for more
} details.
}
} Second, although I've had very little to do with TEMs over the years, most
} typically run at much lower emission currents than SEMs, in the range of
} 10-15�A, I believe. �What does your user's manual recommend?
}
} The emission current could be the whole answer to your short filament life.
}
} Ken Converse
} owner
}
} QUALITY IMAGES
} Servicing Scanning Electron Microscopes
} Since 1981
} 474 So. Bridgton Rd.
} Bridgton, ME �04009
} 207-647-4348
} kenconverse-at-qualityimages.biz
} qualityimages.biz
}
}
} -----Original Message-----
} X-from: one_twinklestar-at-yahoo.com.sg [mailto:one_twinklestar-at-yahoo.com.sg]
} Sent: Tuesday, April 24, 2012 12:43 PM
} To: kenconverse-at-qualityimages.biz
} Subject: [Microscopy] Lab6 Filament Problems
}
}
}
}
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}
} Dear All, a very good day to you! My lab owns a JEOL 2010 TEM. Recently, the
} JEOL engineers have come to do a preventive�maintenance�for our system. They
} have change a new LaB6 filament and done a thorough check on any possible
} leakage and the power stability because before the�maintenance, our filament
} doesn't seems to be running in good condition. The filament current (we set
} at 104uA to 106 uA) drops�noticeably�faster to the background during usage
} and finally burn off (in a span of 4 months). While i am hoping that
} the�maintenance�would clean up the column and stop any possible leakage
} (they did detect a leak at V2, which is at around the camera chamber), the
} new filament which we are using this time doesn't look promising either.
}
} Firstly, before we allow the users to use (when tem is not in use), we
} observed that the filament is unstable �twice. Once is when the current
} suddenly drops from 101uA to zero when not in use. The second time is when
} the JEOL engineer is removing the sample holder (he is suppose to be an
} expert in his own machine). For the second case, the engineer told me that
} it is not because of any discharge from the filament by observing the change
} in the SIP.�Just from yesterday till today, the filament current drops from
} 106 uA (the engineer set at this current) back to 101uA �after a day of
} usage (many users came in to use this TEM). On the other hand, i try to
} observe the shape of the LaB6 filament under undersaturated condition. I did
} see 4 lines from different directions connecting to a dot (supposedly�it
} should be the tip, please correct me if i am wrong), but they seems to be
} tilted at an angle as the dot is not at the center of the beam observed
} under
} �the viewing screen. I try to use the gun tilt in attempt to bring the
} filament back to the center of the beam but it didn't seems to be working.
} What i observed is that the lines and the dots are fading fast into the beam
} (which also darkens) when i do the gun tilt.
}
} I would like to ask
}
} 1) If anyone has experience the fast dropping beam current during usage, if
} so what could be the possible reason
} 2) If i cannot bring the tip back to the center when using the gun tilt,
} what could have happen or what other alignment i could have use?
}
} Pardon me because i am still learning how to use a TEM correctly.
}
} Best Regards,
} Yee Yan, Tay
} FACTS Lab
} Singapore
}
}
} ==============================Original Headers==============================
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

The topic I would like feedback on is how do institutions handle pilot project funding for grant submissions? We waive fees for some of this work, but can't accommodate it all and administration would like to set up a formal structure to support this.

So the questions would be:
Does your institution do this?
What is the amount of money budgeted?
What types of restrictions/guidelines are in place?
Is there follow up on grant submission success rates?

Feel free to mention other topics I might have overlooked.

Thanks in advance!

Randy

Randy Nessler
Director
Central Microscopy Research Facility
University of Iowa
Phone 319-335-8142

________________________________
Notice: This UI Health Care e-mail (including attachments) is covered by the Electronic Communications Privacy Act, 18 U.S.C. 2510-2521, is confidential and may be legally privileged. If you are not the intended recipient, you are hereby notified that any retention, dissemination, distribution, or copying of this communication is strictly prohibited. Please reply to the sender that you have received the message in error, then delete it. Thank you.
________________________________

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12, 27 -- From randy-nessler-at-uiowa.edu Thu May 3 10:23:52 2012
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12, 27 -- From: "Nessler, Randy A" {randy-nessler-at-uiowa.edu}
12, 27 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
12, 27 -- Subject: Funding pilot projects
12, 27 -- Thread-Topic: Funding pilot projects
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Contents Retrieved from Microscopy Listserver Archives
http://www.microscopylistserver.com/MicroscopyListserver/MicroscopyArchives.html

Here at the University of Tulsa there is an internal research grant system that basically allows for preliminary investigations.
It isn't specifically for microscope fees, but can be used for them.
Amounts are up to $1500 per semester for faculty, $500 for students, and re-applications are allowed.
Our internal rates are about $50 per hour, so generally there's plenty to see if a project is viable.

This is all handled by our research office; they review the short (two-page) grant applications, the equally short reports, as well as any follow-ups.
It's a great system for the lab because it's largely invisible (though I am sometimes consulted for comment), and I don't have the burden of deciding who gets the money or whether it's being used appropriately.

Paige Johnson
Nanocharacterization and Fabrication Laboratory
University of Tulsa

-----Original Message-----
X-from: randy-nessler-at-uiowa.edu [mailto:randy-nessler-at-uiowa.edu]
Sent: Thursday, May 03, 2012 10:34 AM
To: Johnson, Paige

The topic I would like feedback on is how do institutions handle pilot project funding for grant submissions? We waive fees for some of this work, but can't accommodate it all and administration would like to set up a formal structure to support this.

So the questions would be:
Does your institution do this?
What is the amount of money budgeted?
What types of restrictions/guidelines are in place?
Is there follow up on grant submission success rates?

Feel free to mention other topics I might have overlooked.

Thanks in advance!

Randy

Randy Nessler
Director
Central Microscopy Research Facility
University of Iowa
Phone 319-335-8142

________________________________
Notice: This UI Health Care e-mail (including attachments) is covered by the Electronic Communications Privacy Act, 18 U.S.C. 2510-2521, is confidential and may be legally privileged. If you are not the intended recipient, you are hereby notified that any retention, dissemination, distribution, or copying of this communication is strictly prohibited. Please reply to the sender that you have received the message in error, then delete it. Thank you.
________________________________

==============================Original Headers==============================
12, 27 -- From randy-nessler-at-uiowa.edu Thu May 3 10:23:52 2012
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12, 27 -- From: "Nessler, Randy A" {randy-nessler-at-uiowa.edu}
12, 27 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
12, 27 -- Subject: Funding pilot projects
12, 27 -- Thread-Topic: Funding pilot projects
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I'm helping a friend set up his SEM and EDS at his venture. We are locked in a titanic struggle over working distance and take-off angle. (Well, it's not that big and we can both live with the other's suggestion. No SiLi crystals have been harmed...) He has an windowless EDS detector on a SEM, what take-off angle and working distance should he use?

I'm avoiding stating our positions but what does the collective wisdom or the microscopy community say?

Frank

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Hi Debby,
We ordered a unit from GEditigal energy (www.gedigitalenergy.com); different specifications than yours (5kvA) but we've been very pleased.
And when we placed the order they gave us a bigger educational discount than we had expected!

Paige Johnson
Nanocharacterization and Fabrication Laboratory
University of Tulsa

-----Original Message-----
X-from: dsherman-at-purdue.edu [mailto:dsherman-at-purdue.edu]
Sent: Tuesday, May 01, 2012 7:23 PM
To: Johnson, Paige

Hi all,

I want to purchase a UPS system for our SEMs. Would appreciate information
as to vendors. Specifications we need are:
- Rating = 8 kVA / 6.4 kW
- Typical backup time of core instrument (i.e. at ca. 27% load) = 60 min.
- Input voltage = 172 - 285 Vac
- Input frequency = 40 - 70 Hz
- Output voltage = 230 Vac
- Output frequency = 50 / 60 Hz.

Any suggestions of vendors or internet sites that may have appropriate
systems would be appreciated.

Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

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17, 31 -- From paige-johnson-at-utulsa.edu Thu May 3 11:07:01 2012
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I thought the two were pretty much determined by the adapter plate for the EDS. There may be some leeway in the design of that plate.

If the detector is already mounted, I would simply check the count rate as a function of working distance. The collimator on the EDS detector will restrict you to some range. I think the proper distance would be where you get the maximum count rate. You would then need to determine the take-off angle and enter that with the EDS parameters. The collimator may be cutout at the bottom to let you use longer distances at less-than-optimum intensity. The working distance and take-off angle should be reentered for those spectra for the best quant results.

If the plate has not yet been made, my general impression was that it was best to design the system for the largest solid angle for the detector. The solid angle (and count rate) will go as one over the square of the distance, so a little change will make a big difference. I am always looking for more count rate so I can keep my beam as small as possible.

The counter argument is that you want to have a short working distance to be able to do EDS at the same working distance that is used for the best imaging. It is a relatively small matter to change the condenser lens to increase the current. It is less convenient to pull the EDS detector back, raise the sample, and change the current.

I would tend to weigh solid-angle as the most important factor, but I would probably insist on a take-off angle of 30 degrees or more. I would probably accept a longer working distance, if necessary, since imaging performance seems to fall off less quickly with distance than does x-ray count rate.

The question may come down to what the priority will be for the system. We have an older W-gun system that is used for a lot of EDS. It has an analytical working distance of 25 mm. We need lots of beam to get a decent count rate so we know we are going to be sacrificing spatial resolution. We pull the detector back, raise the sample, and choke down the beam for better imaging.

We have a newer FE-SEM that is used primarily for imaging. It is normally run at 10-mm working distance. For critical samples, we can shorten that distance. But it also has a sharper pole piece so that EDS can be done at 10 mm. We only need to change the beam current.

I'll be interested in hearing the opposite sides of your debate.

Warren

-----Original Message-----
X-from: frank_karl-at-ardl.com [mailto:frank_karl-at-ardl.com]
Sent: Thursday, May 03, 2012 11:07 AM
To: wesaia-at-iastate.edu

I'm helping a friend set up his SEM and EDS at his venture. We are locked in a titanic struggle over working distance and take-off angle. (Well, it's not that big and we can both live with the other's suggestion. No SiLi crystals have been harmed...) He has an windowless EDS detector on a SEM, what take-off angle and working distance should he use?

I'm avoiding stating our positions but what does the collective wisdom or the microscopy community say?

Frank

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Hi Frank

So often when I visit a client the x-ray set up is at the generic value that
the service technician has given to them; I do not believe it! Simply the
slop in the fixing bolts and minor constructional changes will change the
"perfect" x-ray position. All of these instruments have character, no two
from the same manufacturer are exactly the same, you always need to search
for the "perfect" x-ray working distance.

How should you fix the problem? Well you need to find the WD that gives the
highest x-ray signal as only in that position will the most trace element
data be visible. So place a naked stub in the system and at a fixed
magnification run a spectrum and note the count rate. Move to another level
and check the spectrum again at the same magnification and having corrected
the focus. Repeat until within plus or minus 1mm the best count rates are
achieved.

On a windowless system, with the bulky head, I would start at a very long
WD, say 45mm, and work slowly upwards towards 25mm but I think that would be
too short in my experience.

Good luck it's not such a problem.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: frank_karl-at-ardl.com [mailto:frank_karl-at-ardl.com]
Sent: 03 May 2012 17:07
To: protrain-at-emcourses.com

I'm helping a friend set up his SEM and EDS at his venture. We are locked
in a titanic struggle over working distance and take-off angle. (Well, it's
not that big and we can both live with the other's suggestion. No SiLi
crystals have been harmed...) He has an windowless EDS detector on a SEM,
what take-off angle and working distance should he use?

I'm avoiding stating our positions but what does the collective wisdom or
the microscopy community say?

Frank

This email and any of its attachments may contain confidential information
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received this email in error, please notify us by return email at
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this email contains test data and/or draft reports, you are hereby notified
that only a signed original test report will contain official results, a
copy of which resides in the project folder located at ARDL, Inc. Thank you.
Akron Rubber Development Laboratory, Inc.

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On May 2, 2012, at 1:45 AM, stefan.diller-at-t-online.de wrote:

} is there an easy way to make a gold on carbon test specimen for SEM ?

Dear Stefan,
If you just want gold islands on a carbon surface, you can evaporate
carbon onto a formvar-covered grid, then briefly evaporate a small
amount of gold onto that, but if you want both islands of gold and
other islands of graphite, you will need to deposit the carbon in such
a way that the graphite islands form, and I do not know the procedure
for that. It may be more cost- and time-effective to buy a combined
test specimen from one of the supply dealers, which should be about
$50 or somewhat less.
Yours,
Bill

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Hi Randy

We have just introduced what we are calling our Zero Fees Schemme. Details at

http://ocem.otago.ac.nz/zero_fee_scheme.html

Happy to answer any questions

Have a nice day

Allan

On 4/05/2012, at 3:31 AM, {randy-nessler-at-uiowa.edu} {randy-nessler-at-uiowa.edu} wrote:

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} The topic I would like feedback on is how do institutions handle pilot project funding for grant submissions? We waive fees for some of this work, but can't accommodate it all and administration would like to set up a formal structure to support this.
}
} So the questions would be:
} Does your institution do this?
} What is the amount of money budgeted?
} What types of restrictions/guidelines are in place?
} Is there follow up on grant submission success rates?
}
} Feel free to mention other topics I might have overlooked.
}
} Thanks in advance!
}
} Randy
}
} Randy Nessler
} Director
} Central Microscopy Research Facility
} University of Iowa
} Phone 319-335-8142
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} 12, 27 -- From randy-nessler-at-uiowa.edu Thu May 3 10:23:52 2012
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11, 23 -- From allan.mitchell-at-stonebow.otago.ac.nz Thu May 3 15:27:46 2012
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Allan Mitchell posted an entry in this thread referring to their "Zero
Fees Scheme". It sounds intelligent and wonderful. However, they are
in New Zealand, and laboratories in the USA should remember that they
may not offer a scheme like this if their instruments have Federal
funding. In the USA, research on Federal funding must be offered the
instruments at the lowest rate. Therefore, if there is a "zero fees
scheme" all federally-funded research would have to be free. Similarly,
if classes are not charged, research would have to be free. There was a
thread on this recently.

Alwyn Eades

Department of Materials Science and Engineering
Lehigh University

==============================Original Headers==============================
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I am not sure I agree with this interpretation of the federal rules. They are essentially awarding internal grants. The core simply has to "bill" the funding agency (e.g., the university's office of research) the same rate as they bill federally funded users. If it was free to anyone for as much as they want unless they had a grant, that would be a violation. If the school gives a "grant" of 20 hrs of confocal time, how is that different from giving a start up package to a new Asst Prof that is used to pay for core fees. Assuming a core is not 100% self-sufficient, it is simply a different way to deliver a subsidy to the core.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: jae5-at-lehigh.edu [mailto:jae5-at-lehigh.edu]
Sent: Thursday, May 03, 2012 3:45 PM
To: Phillips, Thomas E.

Allan Mitchell posted an entry in this thread referring to their "Zero Fees Scheme". It sounds intelligent and wonderful. However, they are in New Zealand, and laboratories in the USA should remember that they may not offer a scheme like this if their instruments have Federal funding. In the USA, research on Federal funding must be offered the instruments at the lowest rate. Therefore, if there is a "zero fees scheme" all federally-funded research would have to be free. Similarly, if classes are not charged, research would have to be free. There was a thread on this recently.

Alwyn Eades

Department of Materials Science and Engineering Lehigh University

==============================Original Headers==============================
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15, 27 -- From PhillipsT-at-missouri.edu Thu May 3 17:25:13 2012
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We have a pilot project fee of $25 for sem work no matter who requires it.
Therefore it is the lowest fee for a specific type of work. We did
consider a no-fee but our time is involved, so a small amount was
necessary and you usually know within half to one hour on the microscope
whether the project will work.
Elaine

Dr. Elaine C. Humphrey
STEHM Technologist and Lab Manager
Bob Wright Science Centre A015
Advanced Microscopy Facility
University of Victoria, Canada

Lab: 250-853-3968
cell: 250-886-2068
website: http://www.stehm.uvic.ca

On 03/05/12 3:33 PM, "PhillipsT-at-missouri.edu" {PhillipsT-at-missouri.edu}
wrote:

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12, 32 -- From ech-at-uvic.ca Thu May 3 17:57:18 2012
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Thanks to all for the suggestions and other information regarding UPS systems to protect our major instruments. I suggest that you consider including a UPS system in all new EM purchases. It is usually easier to roll the price into the original instrument pricing than try to come u with the funds at a later date.

Debby

=====================================================

Hi all,

I want to purchase a UPS system for our SEMs. Would appreciate information
as to vendors. Specifications we need are:
- Rating = 8 kVA / 6.4 kW
- Typical backup time of core instrument (i.e. at ca. 27% load) = 60 min.
- Input voltage = 172 - 285 Vac
- Input frequency = 40 - 70 Hz
- Output voltage = 230 Vac
- Output frequency = 50 / 60 Hz.

Any suggestions of vendors or internet sites that may have appropriate
systems would be appreciated.

Debby

---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896
Purdue University E-mail: dsherman-at-purdue.edu
S-052 Whistler Building
170 S. University Street
West Lafayette, IN 47907
http://www.ag.purdue.edu/facilities/microscopy

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Dear Stefan, dear all,

not dealing with such problems during my work but interested in matters like yours:

seconding what Bill said, I would like to point you to an open paper.pdf you perhaps haven't I your collection:

http://www.osti.gov/bridge/servlets/purl/95473-5ReeGR/webviewable/95473.pdf

LEE RH, 1995 (Argonne Natl. Labs) (I think that's where Nestor is working)
{"Coatings and Alternatives for SEM Microscopy" (Presented at INTER/MICR0'93, July 19, 1993, Chicago, IL)

perhaps you'll find an answer (cf. lower p.3) (==} island formation perhaps "automatically"? due to the {tendency} of gold to form islands on carbon)

"The main disadvantage of gold and silver coatings is their tendency to migrate
on the surface of the sample and coalesce into islands or particles. This property
is utilized to make resolution test samples for the scanning electron microscope (SEM)."

Further/additional finding of "island" in the text: see p 12

On the other hand, there are the supply dealers offering a variety of calibration and test specs.
(e.g. google: Test Specimens and Calibration Standards)

Have a beautiful weekend,
good luck, best wishes and regards

Wolfgang MUSS
SALZBURG-AUSTRIA
[posted on-line also for colleagues/"listers" interested in {historical} lit-refs]

Von: wtivol-at-sbcglobal.net [mailto:wtivol-at-sbcglobal.net]
Gesendet: Donnerstag, 03. Mai 2012 22:03
An: Mu� Wolfgang
Betreff: [Microscopy] Re: Gold on carbon test specimen

----------------------------------------------------------------------------
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On May 2, 2012, at 1:45 AM, stefan.diller-at-t-online.de wrote:

} is there an easy way to make a gold on carbon test specimen for SEM ?

Dear Stefan,
If you just want gold islands on a carbon surface, you can evaporate carbon onto a formvar-covered grid, then briefly evaporate a small amount of gold onto that, but if you want both islands of gold and other islands of graphite, you will need to deposit the carbon in such a way that the graphite islands form, and I do not know the procedure for that.

It may be more cost- and time-effective to buy a combined test specimen from one of the supply dealers, which should be about $50 or somewhat less.
Yours,
Bill

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Hi Frank,
I think Warren and Steve covered the WD and solid angle aspects pretty well.
The remaining question is tilt.

Does your detector enter horizontally or is it angled down? Generally if
it's angled down, the specimen is run level and the best WD is where the
axis of the detector intersects the axis of the beam. Keeping the specimen
horizontal (perpendicular to the beam) gives the smallest interaction volume
for the x-rays. Tilting it gives more counts but generally from a larger
part of the surface due to the beam running at an angle under the surface.

If the detector enters horizontally you have choices to make and I'm not
expert enough to give you all the ins and outs, but here are some
possibilities.

Run the sample horizontally and adjust the WD for the best count rate. This
gives you the smallest interaction volume but the counts may suffer.

Use a 30 deg tilt (or something else that strikes your fancy) and look for
the best WD. You'll get better counts with some loss of spatial resolution.

Use a 45 deg tilt and a WD that puts you on the centerline of the detector.
This may give you your best counts with some more loss of spatial
resolution. On the other hand, moving the detector in and out doesn't
change the take-off angle, only the solid angle, which will give you more
consistent results quantitatively if you plan on moving the detector in and
out.

Now let's hear the 2 arguments from you.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: frank_karl-at-ardl.com [mailto:frank_karl-at-ardl.com]
Sent: Thursday, May 03, 2012 12:08 PM
To: kenconverse-at-qualityimages.biz

I'm helping a friend set up his SEM and EDS at his venture. We are locked
in a titanic struggle over working distance and take-off angle. (Well, it's
not that big and we can both live with the other's suggestion. No SiLi
crystals have been harmed...) He has an windowless EDS detector on a SEM,
what take-off angle and working distance should he use?

I'm avoiding stating our positions but what does the collective wisdom or
the microscopy community say?

Frank

This email and any of its attachments may contain confidential information
intended only for the use of the addressee(s). If the reader of this email
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this email contains test data and/or draft reports, you are hereby notified
that only a signed original test report will contain official results, a
copy of which resides in the project folder located at ARDL, Inc. Thank you.
Akron Rubber Development Laboratory, Inc.

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-------- Original Message --------

-------- Original Message --------

Hi Zack

Pleased I could "count" on making your day, now you are one of the
professionals! I wonder how many others have checked their system out?

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: Zack Gainsforth [mailto:zackg-at-berkeley.edu]
Sent: 05 May 2012 00:41
To: protrain-at-emcourses.com

Greetings Fellow Microscopists,

Two advanced scanning electron microscopy short courses will be instructed by Dr. Craig Schwandt of The McCrone Group at Hooke College of Applied Sciences, located in Westmont, IL this July.

INS-605: ADVANCED X-RAY MICROANALYSIS BY EDS
This is an advanced course in x-ray microanalysis using energy-dispersive x-ray spectrometry (EDS). The course will provide instruction and hands-on practice on performing EDS, analyzing difficult samples, mapping, and interpreting analysis results.
http://www.hookecollege.com/courses/course.asp?COURSE_ID=129

INS-610: ADVANCED IMAGING TECHNIQUES FOR SCANNING ELECTRON MICROSCOPY
This Scanning Electron Microscopy (SEM) imaging course is an advanced topics course and provides instruction and hands-on practice for getting the best possible SEM images, especially from difficult samples, or under challenging operating conditions. Signals and image generation, instrument operation, operating variables, image interpretation and applications of SEM will be studied through lectures and hands-on activities. Advanced SEM imaging topics such as very high resolution imaging and low voltage imaging will be covered.
http://www.hookecollege.com/courses/course.asp?COURSE_ID=127

Best regards-

__________________________________________________
Chris Gorman
Admissions Specialist
Hooke College of Applied Sciences
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7412(fax)
www.hookecollege.com

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We have a great opportunity for an experienced electron/light
microscopist to work in a diverse and energetic core facility. The
benefits include the opportunity to design and implement a wide
variety of experimental procedures, to work with a varied group of
investigators, to operate state of the art equipment and be part of a
constructive and resourceful team. This position is in the Johns
Hopkins School of Medicine Microscope Facility (link at bottom of message).

Description: We seek a microscopy specialist talented in electron
microscopy specimen preparation, particularly thin
sectioning. Additional fluorescence/confocal microscopy is
advantageous. Not only helping to develop techniques to bridge the
two microscope modalities, the candidate will help assist in training
and supervising new users, as well as help users troubleshoot
experiments and assist in sample preparation as needed. Strong
analytical and organizational skills coupled with strong
interpersonal and communication skills (both oral and written) are essential.

Environment: With 4 other staff members, the candidate will provide
user support at the Johns Hopkins School of Medicine Microscope
Facility. This facility provides light, fluorescence and electron
microscopy services to 350+ users throughout Johns Hopkins using 17+
advanced microscopes and sophisticated preparatory equipment. Because
of the diversity of equipment within the facility, the candidate must
display resourceful independence and a willingness to learn.
Ultimately, the candidate is expected to become an expert resource
for image quantitation, as well as a resource for using, analyzing,
and troubleshooting experiments with fluorescence/confocal microscopy
spanning generic and highly specialized applications.

Duties: The primary duties of the candidate will be to provide
microscopy services, both in electron microscopy and in
fluorescence/confocal microscopy. Basic knowledge of cell biology is
critical for communicating with users. Helping users with specimen
preparation and interpretation may be needed as part of user services
on projects. Regular duties include cleaning these microscope work
areas and maintaining relevant supplies. Secondary duties will
include collecting and maintaining a library of protocols, manuals
and tutorials for users. With all duties, timely record-keeping
(electronic & written) are required.

Posted Qualifications:
BS/BA in biological sciences, chemistry, or related field required.
At least three years of relevant work experience is required.

To apply follow the link for job requisition 50957:
https://hrnt.jhu.edu/jhujobs/job_view.cfm?view_req_id=50957&view=sch

=================================================================================
Scot C. Kuo, (410)955-4536, skuo-at-jhu.edu
Director, Microscope Facility & Imaging for IBBS;
www.hopkinsmedicine.org/micfac/
Associate Professor, Biomedical Engineering & Cell Biology; www.jhu.edu/cmml/
Wood Basic Science Bldg, Room G10, 725 N Wolfe St, Baltimore MD 21205
=================================================================================

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Hello Listservers,

We have been doing HPF/AFS for over a year on various samples, and have had great success with membrane morphology. Just recently we have run into a snag, our drosophila embryos do not show any membranes!!! The samples are frozen in fresh yeast paste (as before when it worked), stored and transferred to frozen cocktails (1% osmium (acetone) 0.1% Uranyl Acetate (MEOH) 5% D-H20) under liquid nitrogen and processed for AFS as normal. The question is why the membrane loss? Could it be microgas pockets in the yeast paste? Could stored frozen cocktail mixes lose their efficacy? Is it just a bad "Freeze" (although the numbers looked right). Could the yeast paste be too thick? Is this the variability we have to deal with this technique? Any thoughts will be greatly appreciated.

sincerely,

Michael Delannoy

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Nester and fellow Microscopists,
I am sending this email through Firefox to check whether Explorer was the culprit in all my recent rejected emails, sorry for the inconvenience.

Michael Delannoy

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Listers,
Recently I had Company X rebuild a 5010 Alcatel molecular drag
pump(MDP) that was not pulling the vacuum I needed. This MDP serves as
a backing pump to a larger turbo pump. Upon receiving the rebuilt MDP
I placed it back into the vacuum setup but was not able to get the MDP
to accelerate to full capacity, the rebuilt MDP pulls a vacuum to
~1000 Pa. The backing pressure(from a diaphragm pump) is roughly 6750
Pa, a backing pressure that has always been sufficient for "my" 5010
Alcatel MDPs.
Since confirming proper backing pressure I have contacted Company X
stating my issue and they retested the rebuilt MDP, and still state
that the pump pulls a vacuum of 6x10^-6 mbar (6x10^-4 Pa). I have a
second MDP, another 5010 Alcatel MDP, which accelerates properly(
better than 1 Pa) under five minutes when used in the exact same
vacuum setup where the rebuilt MDP will not fully accelerate. To me,
using this second MDP allows the backing pressure and potential leaks
within tubes/etc to be ruled out as a possibility for the rebuilt MDP
not to operate at full capacity, agreed? Are there other options for
me to test whether the rebuilt MDP is operating correctly? Am I able
to lean harder on Company X when I know a virtually identical MDP
performs properly within in my vacuum setup and when I replace the
functioning MDP for the rebuilt MDP and the result is basically
failure? Any suggestions or thoughts on this matter will be greatly
appreciated.

-Joe

--
Joseph Heintz
Lab/BBPIC Manager
University of Wisconsin-Madison
Room 1048, Animal Sciences Building
1675 Observatory Dr.
Madison, WI 53706
608.263.4162

==============================Original Headers==============================
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Hi,

Typically when I need to get better spatial resolution out of EDS maps, I've been using low voltage, which works quite well. I commonly see 100 or 200 nm resolutions and can still quantify most elements. In part my choice to use low voltage is due to the fact that my samples tend to be a bit delicate, but recently I've been doing some EDS on an iron meteorite, and it can take a bit more of a pounding than my usual sample. So, I decided to give grazing exit a spin and it really works nicely. I get about 100 nm resolution on my tungsten SEM at 20 keV, and the count rates are roughly comparable to using low voltage with my setup (at least for this material).

So my question is this: Do any of you have much experience with the specific strengths and weaknesses of these two methods? As I see it so far, I'm noticing that quantitation with grazing exit is tough because it is so dependent on the exit angle and solid angle. But it gives good count rates, good spatial resolution, and allows me to use to higher energy k-shells so it is easier to see (or separate) many elements. Low voltage is typically less finicky to set up and gives much better quantitative results (at least for me). Grazing exit also seems to be useful only on samples that can take a fair bit of beating because it takes a little while to get the sample and detector oriented into the evanscecent regime (or at least close to it).

Does anyone else have experience/thoughts on this?

Zack Gainsforth
Space Sciences Laboratory, UC Berkeley
7 Gauss Way
Berkeley, CA 94720
510-642-9733
zackg-at-ssl.berkeley.edu

Zack

==============================Original Headers==============================
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Joe,

When you said that you used the same vacuum setup when testing the second
pump, did that mean that you used the same controller as the first pump? If
they are testing the pump and it gets up to speed at their test facility,
then the pump is probably OK. You have only a couple of potential problems.

1) You have a problem with the controller. It also could be a problem with
connectors and cabling. Since the pump worked during testing at company X,
the connector on the pump is probably OK. Check your cables and your cable
connectors.
2) you have a vacuum leak. Anytime you make or break a vacuum connection,
you have the potential for a leak. Look at your O-rings and your sealing
surfaces for scratches on that first pump.
3) you have a problem with the backing pump. Since you tested that with the
second pump, this shouldn't be a problem.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: jheintz-at-wisc.edu [mailto:jheintz-at-wisc.edu]
Sent: Tuesday, May 08, 2012 2:11 PM
To: s.walck-at-cox.net

Listers,
Recently I had Company X rebuild a 5010 Alcatel molecular drag
pump(MDP) that was not pulling the vacuum I needed. This MDP serves as a
backing pump to a larger turbo pump. Upon receiving the rebuilt MDP I placed
it back into the vacuum setup but was not able to get the MDP to accelerate
to full capacity, the rebuilt MDP pulls a vacuum to
~1000 Pa. The backing pressure(from a diaphragm pump) is roughly 6750 Pa, a
backing pressure that has always been sufficient for "my" 5010 Alcatel MDPs.
Since confirming proper backing pressure I have contacted Company X
stating my issue and they retested the rebuilt MDP, and still state that the
pump pulls a vacuum of 6x10^-6 mbar (6x10^-4 Pa). I have a second MDP,
another 5010 Alcatel MDP, which accelerates properly( better than 1 Pa)
under five minutes when used in the exact same vacuum setup where the
rebuilt MDP will not fully accelerate. To me, using this second MDP allows
the backing pressure and potential leaks within tubes/etc to be ruled out as
a possibility for the rebuilt MDP not to operate at full capacity, agreed?
Are there other options for me to test whether the rebuilt MDP is operating
correctly? Am I able to lean harder on Company X when I know a virtually
identical MDP performs properly within in my vacuum setup and when I replace
the functioning MDP for the rebuilt MDP and the result is basically failure?
Any suggestions or thoughts on this matter will be greatly appreciated.

-Joe

--
Joseph Heintz
Lab/BBPIC Manager
University of Wisconsin-Madison
Room 1048, Animal Sciences Building
1675 Observatory Dr.
Madison, WI 53706
608.263.4162

==============================Original Headers==============================
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Email: mgengle-at-uky.edu
Name: Mary Gail Engle

Organization: University of KY

Title-Subject: service contract providers

Message: Liz,
Several years ago we were also required to abandon our service contracts to go with a provider
(Thermo) and of course we all went kicking and screaming the whole way. I had visions of the local
electrician coming in to work on our EM and the laser lightshow guy repairing our confocals.
However, it's worked out very well because they let us use our manufacturers (Leica and FEI) and
therefore we get the service people we've always used. I'm not so sure the microscope companies
love the program as they can't plan budgets too well with all of us calling only when we need
them... But we do get service, albeit more slowly than if we'd had a contract.

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Email: mdelann1-at-jhmi.edu
Name: Michael Delannoy

Organization: Johns Hopkins School of Medicine Microscope Facility

Title-Subject: Microscopy Specialist Job Opportunity

Message: We have a great opportunity for an experienced electron/light microscopist, to work in a
diverse and energetic core facility. The benefits include the opportunity to design and implement a
wide variety of experimental procedures, to work with a varied group of investigators, to operate
state of the art equipment and be part of a constructive and resourceful team. This position is in
the Johns Hopkins School of Medicine Microscope Facility (est. 1989).

Description: We seek a microscopy specialist talented in electron microscopy specimen preparation,
particularly thin sectioning. Additional fluorescence/confocal microscopy is advantageous. Not
only helping to develop techniques to bridge the two microscope modalities, the candidate will help
assist in training and supervising new users, as well as help users troubleshoot experiments and
assist in sample preparation as needed. Strong analytical and organizational skills coupled with
strong interpersonal and communication skills (both oral and written) are essential.

Environment: With 4 other staff members, the candidate will provide user support at the Johns
Hopkins School of Medicine Microscope Facility. This facility provides light, fluorescence and
electron microscopy services to 350+ users throughout Johns Hopkins using 17+ advanced microscopes
and sophisticated preparatory equipment. Because of the diversity of equipment within the facility,
the candidate must display resourceful independence and a willingness to learn. Ultimately, the
candidate is expected to become an expert resource for image quantitation, as well as a resource for
using, analyzing, and troubleshooting experiments with fluorescence/confocal microscopy spanning
generic and highly specialized applications.

Duties: The primary duties of the candidate will be to provide microscopy services, both in electron
microscopy and in fluorescence/confocal microscopy. Basic knowledge of cell biology is critical for
communicating with users. Helping users with specimen preparation and interpretation may be needed
as part of user services on projects. Regular duties include cleaning these microscope work areas
and maintaining relevant supplies. Secondary duties will include collecting and maintaining a
library of protocols, manuals and tutorials for users. With all duties, timely record-keeping
(electronic & written) are required.

Posted Qualifications:
BS/BA in biological sciences, chemistry, or related field required. At least three years of relevant
work experience is required.

To apply follow the link for job requisition 50957:
https://hrnt.jhu.edu/jhujobs/job_view.cfm?view_req_id=50957&view=sch

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Email: edward.jarvis-at-timken.com
Name: Edward Jarvis

Organization: The Timken Company

Title-Subject: Historical Leitz Objective Lens

Message: Listservers

We are in need of an objective lens approximately 156X to 160X for a Leitz Orthoplan microscope. If
any one has one that is no longer needed, please contact me. Edward.Jarvis-at-Timken.com

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} } }
} Nester and fellow Microscopists,
} I am sending this email through Firefox to check whether Explorer was the
} culprit in all my recent rejected emails, sorry for the inconvenience.
}
} Michael Delannoy

Michael,
I do understand, but ... just a comment on this:
usually, it is not simply "the browser", but the exact version of the browser which matters, the interaction with the operating system, and, most importantly, the settings of the browser, which are numerous and often not checked / not set by the users - but used as they came, during installation.
e.g. which version of firefox are YOU using, today? did you continue to count the 'updates'?
and which IE version? which OS? which settings?
It is not necessary to report this to the listserver, but this is to remind everybody that this is a problem, nowadays, for too many 'users', and it will remain so, for the future.
kind regards,
Reinhard

--
Prof. Dr. Reinhard Rachel
University of Regensburg
Centre for EM / Anatomy
Faculty of Biology & Preclin. Med.
Universitaetsstrasse 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-regensburg.de
office: VKL 3.1.29

next microscopy conferences:
http://www.emc2012.org.uk/
EMC 2012 in Manchester, UK
http://www.mc2013.de/
MC2013 in Regensburg, Germany

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Joe,

I assume you just put pump 2 (good one) in the same position/tubes,
etc. as pump 1 (bad one). Did you also switch controllers? There are
2 controller boxes, although one may be stuffed away somewhere. (I
think it failed in the past.)
If the bad pump is still bad when hooked to the same controller as
the good pump, then you've got a case.

Also, how's the turbo running when hooked up to the bad pump? Speed
OK, sounds good?

Phil
P.S. How about the gauge you're using? You tried it on both pumps,
yes? If only on the bad pump, see how the gauge reads on the good
pump.

} Listers,
} Recently I had Company X rebuild a 5010 Alcatel molecular drag
} pump(MDP) that was not pulling the vacuum I needed. This MDP serves as
} a backing pump to a larger turbo pump. Upon receiving the rebuilt MDP
} I placed it back into the vacuum setup but was not able to get the MDP
} to accelerate to full capacity, the rebuilt MDP pulls a vacuum to
} ~1000 Pa. The backing pressure(from a diaphragm pump) is roughly 6750
} Pa, a backing pressure that has always been sufficient for "my" 5010
} Alcatel MDPs.
} Since confirming proper backing pressure I have contacted Company X
} stating my issue and they retested the rebuilt MDP, and still state
} that the pump pulls a vacuum of 6x10^-6 mbar (6x10^-4 Pa). I have a
} second MDP, another 5010 Alcatel MDP, which accelerates properly(
} better than 1 Pa) under five minutes when used in the exact same
} vacuum setup where the rebuilt MDP will not fully accelerate. To me,
} using this second MDP allows the backing pressure and potential leaks
} within tubes/etc to be ruled out as a possibility for the rebuilt MDP
} not to operate at full capacity, agreed? Are there other options for
} me to test whether the rebuilt MDP is operating correctly? Am I able
} to lean harder on Company X when I know a virtually identical MDP
} performs properly within in my vacuum setup and when I replace the
} functioning MDP for the rebuilt MDP and the result is basically
} failure? Any suggestions or thoughts on this matter will be greatly
} appreciated.
}
} -Joe
}
} --
} Joseph Heintz
} Lab/BBPIC Manager
} University of Wisconsin-Madison
} Room 1048, Animal Sciences Building
} 1675 Observatory Dr.
} Madison, WI 53706
} 608.263.4162

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Hello,

We've received a full suite of standards from the Smithsonian and I'm looking for someone, or a company that would be able to mount them for microprobe work.

Does anyone have any suggestions?

Cheers

Nick

Nicholas J. Foster
SENCR-MIC Laboratory Technician
Fayetteville State University
Office: (910) 672-2039
Probe Lab: (910) 672-2037
Email: nfoster1-at-uncfsu.edu

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Are you sure that Explorer was sending the messages as plain text? I've had messages rejected occasionally when I accidently switched over to a fancier mode.

I don't mind being your guinea pig to test the message format. You can send me a message as if you were sending it to the list. For that matter, copy me on a message that you send to the list and which gets bounced. Then I would know for sure there is something to find.

Warren

-----Original Message-----
X-from: delannoy-at-jhmi.edu [mailto:delannoy-at-jhmi.edu]
Sent: Tuesday, May 08, 2012 2:07 PM
To: wesaia-at-iastate.edu

Nester and fellow Microscopists,
I am sending this email through Firefox to check whether Explorer was the culprit in all my recent rejected emails, sorry for the inconvenience.

Michael Delannoy

==============================Original Headers==============================
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X-from: helmut {mocleica-at-aol.com}

Hello !to my knowledge there is no 160 X objective -this 160 is the tube lenght you should read this
objective mag again it is probably a 100x(160 tubelenght ) Objective Oil

Helmut Patzig

==============================Original Headers==============================
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Dear Nick,

There are a few companies who still mount standards, the two whom I am most familiar with are SPI and Electron Microscopy Science, I have also used Vancouver Petrographic for my thin sections, but have not discussed mounting standards with them. SPI and EMS both have standards available as well and have different mount configurations available, however, SPI will take a long time to complete any mount they do not have on the shelf, EMS I believe will respond more quickly, both produce good results. This service and standard availability are becoming more difficult to have done, I would get everything you can get while supplies last.

Best regards
Tom

-----Original Message-----
X-from: nfoster1-at-uncfsu.edu [mailto:nfoster1-at-uncfsu.edu]
Sent: Wednesday, May 09, 2012 10:52 AM
To: Thomas Beasley

Hello,

We've received a full suite of standards from the Smithsonian and I'm looking for someone, or a company that would be able to mount them for microprobe work.

Does anyone have any suggestions?

Cheers

Nick

Nicholas J. Foster
SENCR-MIC Laboratory Technician
Fayetteville State University
Office: (910) 672-2039
Probe Lab: (910) 672-2037
Email: nfoster1-at-uncfsu.edu

==============================Original Headers==============================
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19, 25 -- From beasley-at-fiu.edu Wed May 9 10:26:57 2012
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19, 25 -- May 2012 11:25:36 -0400
19, 25 -- From: Thomas Beasley {beasley-at-fiu.edu}
19, 25 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
19, 25 -- Subject: RE: [Microscopy] Mircroprobe standard mounting
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Hi Liz,

I agree with Russ, it's best to avoid Remi if possible. We were considering this as an option for our instruments but were told directly by FEI that our service would suffer. Customers with FEI service contracts are always a priority and we would be at the bottom of the list resulting in significant delays in service. As a core facility, we could not afford 9 weeks of downtime. When I mentioned this issue to the Remi Group representatives, they claimed to have satisfied customers that had FEI instruments covered by REMI. When I asked for customer referrals specifically from FEI subscribers, I did not receive them.

Another core facility with the University contracted with Remi for their instruments (primarily Bruker) and did not have issues with delayed service so delay issues may actually be manufacturer specific. However, Remi denied payment for portions of the service invoice from Bruker that had never been an issue previously with either the manufacturer service contract or the ThermoFisher Equipment Management program. This resulted in lengthy exchanges with administration and Remi Group regarding what should be covered. Remi Group also said they would include 2 PM visits per year and that they would coordinate these visits. This was not done by Remi Group and they would not provide a refund for this missed service. In the end, the facility has chosen not to renew their Remi contract because of these issues.

Hope this helps,

Sara
**************
Sara Cole, Ph.D.
Senior Microscopist
Campus Microscopy and Imaging Facility (CMIF)
The Ohio State University
245 Biomedical Research Tower
460 West 12th Avenue
Columbus, OH 43210-1239

Phone 614-292-9786
FAX�� 614-247-8849
WEB�� http://www.cmif.osu.edu

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Monday, May 07, 2012 7:23 PM
To: Cole, Sara

Hi all,

Another aspect of using insurance companies (Remi and others) is to find
out exactly what they will cover. Will they only cover the instrument
being down? We sometimes have an instrument with performance issues.
It is working but not as good as it should. This can become a gray area
for maintenance since the microscope is not "dead in the water" but
unusable for your purposes. We have had great response from our
manufacturer. I have great doubts if the insurance company would invest
the same resources.

Cheers,
Henk

At 5/10/2012 2:46 PM, cole.553-at-osu.edu wrote:
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe --http://www.microscopy.com/MicroscopyListserver
} On-Line Helphttp://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi Liz,
}
} I agree with Russ, it's best to avoid Remi if possible. We were considering this as an option for our instruments but were told directly by FEI that our service would suffer. Customers with FEI service contracts are always a priority and we would be at the bottom of the list resulting in significant delays in service. As a core facility, we could not afford 9 weeks of downtime. When I mentioned this issue to the Remi Group representatives, they claimed to have satisfied customers that had FEI instruments covered by REMI. When I asked for customer referrals specifically from FEI subscribers, I did not receive them.
}
} Another core facility with the University contracted with Remi for their instruments (primarily Bruker) and did not have issues with delayed service so delay issues may actually be manufacturer specific. However, Remi denied payment for portions of the service invoice from Bruker that had never been an issue previously with either the manufacturer service contract or the ThermoFisher Equipment Management program. This resulted in lengthy exchanges with administration and Remi Group regarding what should be covered. Remi Group also said they would include 2 PM visits per year and that they would coordinate these visits. This was not done by Remi Group and they would not provide a refund for this missed service. In the end, the facility has chosen not to renew their Remi contract because of these issues.
}
} Hope this helps,
}
} Sara
} **************
} Sara Cole, Ph.D.
} Senior Microscopist
} Campus Microscopy and Imaging Facility (CMIF)
} The Ohio State University
} 245 Biomedical Research Tower
} 460 West 12th Avenue
} Columbus, OH 43210-1239
}
} Phone 614-292-9786
} FAX 614-247-8849
} WEBhttp://www.cmif.osu.edu
}
}
}
}
}
} -----Original Message-----
} X-from:microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: Monday, May 07, 2012 7:23 PM
} To: Cole, Sara
} Subject: [Microscopy] viaWWW:Remi Group as a equipment service contract provider
}
}
}
}
} ----------------------------------------------------------------------------
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} Date: Mon, 7 May 2012 08:34:03 -0500
} X-from:erwrigh-at-emory.edu ()
}
}
} This Question/Comment was submitted to the Microscopy Listserver using the WWW based Form athttp://www.microscopy.com/MLFormMail.html
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} please copy botherwrigh-at-emory.edu as well as the Microscopy Listserver
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} Email:erwrigh-at-emory.edu
} Name: Elizabeth Wright
}
} Organization: Emory University
}
} Title-Subject: Remi Group as a equipment service contract provider
}
} Message: Dear EM Friends,
}
} Earlier this year, my institution entered into a contract with a company (name available upon
} request) to become the preferred vendor for an Equipment Maintenance Management Program (EMMP). This contract aims to bundle service contracts on equipment in order to decrease university expenses. It was suggested to me that I consider them for servicing our TEMs. I am in direct opposition of this plan for too many reasons to list here.
}
} My question to the list, does anyone have direct, accountable experience with one of these companies that offer this form of service? I would appreciate any and all feedback.
}
} Thank you for your help.
}
} Sincerely,
}
} Liz
}
} Login Host: 170.140.22.142
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
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"Time is that quality of nature which keeps things from happening all at
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Email: gmeliki-at-emory.edu
Name: Gregory Melikian

Organization: Emory University

Title-Subject: Postdoctoral position

Message: A post-doctoral position is available at Emory University in the multidisciplinary laboratory studying the mechanisms of viral entry and fusion with target cells. We employ time-resolved single virus imaging to delineate the mechanisms of entry. The laboratory is equipped with state-of-the-art imaging equipment, including a wide-field deconvolution microscope, a three-color TIRF microscopy system and the Zeiss LSM 780 confocal microscope with FLIM and FCS capabilities. The Emory University provides an excellent collaborative environment and research support through core facilities and common equipment. Highly motivated individuals seeking to develop and apply novel biophysical methodologies to delineate virus entry and fusion are encouraged to apply. Candidates must have a Ph.D. degree in biophysics or related areas. For more information, please visit our website: http://www.pediatrics.emory.edu/divisions/infectious/lab_melikian/index.html. Applicants can contact Dr!
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Email: ravi.thakkar369-at-gmail.com
Name: Ravindra Thakkar

Organization: National Institute of Mental Health and Neuro Sciences

Title-Subject: Common Embedding resing for LM& TEM to carry Medical Diagnosis.

Message: Dear Listener,
We are dealing with the ultra structural pathology of Biopsy samples, So we are seeking for the embedding media which can be used for Light Microscopic as well as for Electron Microscopic studies. Where the semithin sections can be seen in Immunohistochemical studies and its ultrathin section for Transmission Electron Microscopic studies.. Thus we can make able to track cellular information on LM& EM level, and ease of interpretation and save the time also by one time processing..

So kindly suggest for such embedding media and if possible, send me some research article or reports on biopsy work by using such resin.

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Email: watson-at-wi.mit.edu {mailto:watson-at-wi.mit.edu}

Name: Nicki Watson

Organization: whitehead institute

Title-Subject: biotin/avidin label

Message: Dear microscopists,

I am trying to strep Avidin gold label cells that have been biotinalated. W=
e have biotinalated the cells, then fixed with gluteraldehyde 2%, dehydrat=
ed and embedded in plastic. I was under the impression that biotin would su=
rvive this treatment. After sectioning I am labeling with Strep Avidin Gol=
d. I thought I would have a background problem to solve, But I see nothing.

I am wondering if my sample prep is too harsh and I have killed the biotin,=
or if the problem is experimental and the researcher needs to go back to t=
he drawing board.

If any one has any experience post embedding labeling with Avidin-gold plea=
se let me know. The control was to biotinalate, label with avidin and the=
n fix. ( This was also negative.)

Thank you for your input in advance.

Nicki Watson

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{/o:shapelayout} {/xml} {![endif]--} {/head} {body lang=3DEN-US link=3Dblue vli=
nk=3Dpurple} {div class=3DWordSection1} {p class=3DMsoPlainText} Hi Nicki: {o:p=
} {/o:p} {/p} {p class=3DMsoPlainText} You should embed your cells in methacryl=
ates, such as Lowicryl or LR White...n {u} ot epoxy resin {/u} (which does not=
accommodate gold labeling as it is too cross-linked and the surface of the=
epoxy does not swell when the labeling is done). Those resins are us=
ually polymerized at -20C under ultraviolet light. Also, I would use 4.0% p=
araformaldehyde not glut. fixation. {o:p} {/o:p} {/p} {p class=3DMsoPlainText} K=
aren {o:p} {/o:p} {/p} {p class=3DMsoPlainText} {o:p} {/o:p} {/p} {p class=3D=
MsoPlainText} Karen Bentley, M.S. {o:p} {/o:p} {/p} {p class=3DMsoPlainText} Dire=
ctor {o:p} {/o:p} {/p} {p class=3DMsoPlainText} Electron Microscope Research Cor=
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--- {o:p} {/o:p} {/p} {p class=3DMsoPlainText} {o:p} {/o:p} {/p} {p class=3DM=
soPlainText} Email: {a href=3D"mailto:watson-at-wi.mit.edu"} {span style=3D'colo=
r:windowtext;text-decoration:none'} watson-at-wi.mit.edu {/span} {/a} {o:p} {/o:p} {=
/p} {p class=3DMsoPlainText} Name: Nicki Watson {o:p} {/o:p} {/p} {p class=3DMsoP=
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ad institute {o:p} {/o:p} {/p} {p class=3DMsoPlainText} {o:p} {/o:p} {/p} {p =
class=3DMsoPlainText} Title-Subject: biotin/avidin label {o:p} {/o:p} {/p} {p cl=
ass=3DMsoPlainText} {o:p} {/o:p} {/p} {p class=3DMsoPlainText} Message: De=
ar microscopists, {o:p} {/o:p} {/p} {p class=3DMsoPlainText} I am trying to stre=
p Avidin gold label cells that have been biotinalated. We have biotinalated=
the cells, then fixed with gluteraldehyde 2%, dehydrated and embedde=
d in plastic. I was under the impression that biotin would survive this tre=
atment. After sectioning I am labeling with Strep Avidin Gold. I thou=
ght I would have a background problem to solve, But I see nothing. {o:p} {/o:=
p} {/p} {p class=3DMsoPlainText} I am wondering if my sample prep is too harsh=
and I have killed the biotin, or if the problem is experimental and the re=
searcher needs to go back to the drawing board. {o:p} {/o:p} {/p} {p class=3DMs=
oPlainText} If any one has any experience post embedding labeling with Avidi=
n-gold please let me know.   The control was to biotinalate, labe=
l with avidin and then fix. ( This was also negative.) {o:p} {/o:p} {/p} {p cla=
ss=3DMsoPlainText} Thank you for your input in advance. {o:p} {/o:p} {/p} {p cla=
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Hi Fred

Of course there is not "10% of the molarity". That is clear to all.
The formula I suggested using is that if you take 10% of the value of
the molarity of formaldehyde, add it to 30% of the value of the
molarity of glut and add that to the molarity of the buffer, you will
get close to the effective molarity of your fixative.
For example, if you have a 4% solution of formaldehyde, you have a 40g/
L solution or approximately a 1.3M solution. 10% of THAT NUMBER gives
you about 0.13 effective.
It really is very simple, and not at all in need of being told, in
public, that % and molarity are different things.
The person I sent this to understood what I said. Also, several other
people who contacted me understood the formula. Even if you don't
agree with it (it has worked well for me for years), now at least you
understand it.

David

On Apr 12, 2012, at 3:30 PM, Fred Hayes wrote:

} There is no such thing as 10% of the molarity or 30% of the molarity.
} Molarity is expressed as moles/liter or moles of solute/liters of
} solvent. %
} is simply n parts of this vs 100 parts of that. % is concentration
} and has
} nothing to do with molarity. Two different ways of looking at
} solutions
} (solutes/solvent)

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Dear
please, link and upload your abstract at www.owls2012.org! Preliminary program available, list of sponsors in progress.
See you in Genoa soon for a great scientific program.
All the best
Alby

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The last time this happened to me, it was a fried power converter
chip in the power supply - about $250 total to fix. Time before that,
is was a DC/DC power supply chip on the acquisition control board in
the computer. $50 fix.
The other suggestion was a bad chip in the detector pre-amp (but it
wasn't this).

Phil

} Email: mansoreh152-at-gmail.com
} Name: zabihi
}
} Title-Subject: Analysis problem in the SEM
}
} Message: Hi
} The past two weeks I calibrate EDS(Energy Dispersive X-ray
} Spectroscopy)but again, X-ray spectrum are shifted What causes this problem?
} Thanks a lot

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

==============================Original Headers==============================
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Dear All,

Of course, everything is clear, IF, everyone DOES see thru the glass clearly. Unfortunately, everyone does not come to the glass clearly. It always bothered me, especially when instructing students, that when it came to a discussion of fixation, we broke the rules taught them by the chemists and began to use percentages.

So, here's how I began to wrestle with the reality of biologic logic.

1. Buffers. 1% Ca acetate in 4% HCHO --} ~pH 7
4% HCHO = 10% "Formalin" (= ?~37% HCHO, depending on T & P)
HCHO is a gas at ambient T&P, and is VERY soluble in water (special chemistry).
HCHO slowly oxidizes to formic acid - HCOOH (re: buffering)
Formalin left for long periods will show white flecks, milky suspension of an HCHO polymer called paraformaldehyde. HCHO can be regenerated from paraformaldehyde by applying heat and a 'little' base.
HCHO is only rarely described as the 'active' participant in cross-linking.
When I was trained in organic chemistry, I was taught that vic-glycols (vicinal-glycols: a C with two -OH attached functional groups was only a transient (thus, at very small concentrations) component of aldehyde solutions.
However, when a vic-glycol is consumed/withdrawn as a methylene bridge, mass action rules leave 'room' for more vic-glycol to form. (Question: Is this going to be on the test?)(Answer: No! Now I am handing out a one-page protocol that has all of the instructions with none of the 'stuff' that won't be on the test.)

2. Question: What happens to 4% HCHO solutions that must be used routinely in the high Andes medical establishments? Answer: 4% is/was chosen out of habit evolved over years of empirical experience that confirms its functionality as a perfect fixative for most human tissue required for diagnostic purposes. 4% HCHO provides the active cross-linker in great excess. T & P and experience will determine how often fresh 4% needs to be prepared. [Paraffin mixtures - before air conditioning - in the hands of 'expert' histologists were different depending when one was working and the local T, P, and average seasonal relative humidity.]

3. Question: Why do we use a 5-carbon di-aldehyde in TEM preparations. Answer: less shrinkage and more empirical experience!

4. Question: Why can't I use EPON derivatives/'children'/substitutes for large castings? Answer: The published mixtures for TEM are designed to provide 'good' results in small pieces.

5. Question: I have two lots of Methylene Blue. One has a label that claims 35% dye, while the other claims 63%. The protocol calls for 1%. Do I have to do the math? Answer: a) Only if neither works as required by the protocol (Learn about the Biological Stain Commission and how/why certification is used.). b) Assuming a previous working solution of Methylene Blue, compare the equivalent mixtures by applying any of the many relatively 'easy' chemical analysis methods to determine if, indeed, they are all alike. Once you have sufficient data, you MAY be able to apply some math to define a slight change in the protocol to achieve the 'expected' result.

6. Question: How long does the absolute ethanol in your processor work, and what are the relevant ambient conditions there? How about your buddies in Colorado and Florida? Answer: Ask?

7. Question: What are the origins of Wright's stain? Answer: In 1930, H.J. Conn addressed the history of blood stains from which I have extracted the following excerpt. Please take special note of the first sentence in the last of these paragraphs.

----------------
["Nocht (1898) contributed the next important step toward the development of modern blood stains in using Unna's polychrome methylene blue in the Romanovsky combination. Nocht concluded the differential staining obtained by Romanovsky was due in part to decomposition products of methylene blue; and recognizing what Unna had done in the way of intentionally producing these decomposition products, he decided to use the polychrome methylene blue recommended by the latter. First he neutralized the latter solution, using litmus paper to indicate its reaction. He stated that litmus paper could be thus employed by noticing the color in the zone above that stained by the dye itself. Neutralization was accomplished by adding acetic acid until a distinct red appearance was given to litmus paper; this was followed by neutralization with alkali until no red color was observed. This neutralized polychromed solution was then mixed with ordinary methylene blue solution until the violet shade due to the polychroming had disappeared; and was then mixed with eosin. He observed a fine precipitate, but said it was not troublesome.
Nocht did not realize the value of that precipitate; but Jenner (1899) at about the same time collected it and found it a useful stain.
Jenner probably had not seen Nocht's paper and did not polychrome his methylene blue. Instead he mixed ordinary methylene blue with eosin, as Rnmanovsky had done; but instead of staining with the mixture, he filtered off the precipitate formed and redissolved it in methyl alcohol. This methanolic solution he employed as a blood stain in much the manner blood stains are used today. His stain lacked the nuclear staining principle of Romanovsky's and Nocht's stains; but it is still occasionally used today. It is hard to see how anyone today can prefer it to other blood stains; but at the time when it was proposed it was an important step, showing that the staining principle of Romanovsky's stain mas in the precipitate rather than in the part remaining in solution. A practically identical stain was employed by May and Grunwald (1901?), and is in some places called after them rather than after Jenner.
The next step, that of applying Jenner's procedure to the Nocht stain, was such a logical one that it is not surprising to find it occurring to more than one worker simultaneously. Two independent workers, in Germany and England respectively, published papers two years after that of Jenner's in which the procedure was described of collecting and redissolving the Nocht stain precipitate. Reuter (1901) used aniline [auf Deutsch] oil and absolute alcohol as the solvent; Leishmann (1901) dissolved the precipitate in methyl alcohol. Modern blood stains are usually modifications of Leishmann's differing from it only in detail.
In America of recent years the term Wright's stain has come to be almost synonymous with blood stain, to such an extent that stain companies say it is hard to sell one unless it is so labelled. Wright's stain, however, is really a Leishmann stain, differing from the latter only in that the polychrome methylene blue is prepared by heating with sodium bicarbonate for only an hour in flowing steam instead of with sodium carbonate for 1% hours at 65'c, followed by ten days at room temperature. The original popularity of Wright's stain was due to the greater rapidity with which it could be prepared. NOW that biologists buy it already prepared by stain companies, however, they care little whether it is prepared by the method of Leishmann, Wright, or in some other way, as long as it gives the proper results. As a matter of fact, the manufacturers usually do make it by a different procedure; but sell it labelled Wright stain in order to meet the demand.
Stains of the Leishmann-Wright type are thus prepared by starting with indefinite mixtures of simple dyes, and the product is naturally a still more indefinite mixture of compound dyes. Frequent attempts have been made, however, to obtain stains giving similar blood pictures but prepared from compounds of definite chemical composition. It had been recognized quite early by a chemist, Bernthsen (1885), that methylene blue readily yielded certain oxidation products which he called methylene azure and methylene violet. This observation was significant from the standpoint of blood stains, although it was sometime before this significance was appreciated by any biologist. Romanovsky did not realize that his "Rot .aus Methylenblau" had been thus described by a chemist several years earlier. Nocht was familiar with methylene violet; but could not identify the "Rot aus Methylenblau," merely stating that it was neither methylene blue, nor methylene violet.", from H.J. Conn, "History of Staining. THE STAINING OF BLOOD AND PARASITIC PROTOZOA", Stain Technology, 4(5), 127-134, 1930.] [H.J. Conn was a leader in establishing the Certification process for 'standardizing' commonly used/manufactured stains/dyes in order to bring order into their uses - primarily diagnostic.]
------------------------------------
The above could represent the humor in biology, or the truth.
------------------------------------
8. Question: How can the structure of Lactate Dehydrognease (LDH) be utilized in a classroom to teach (hint at) the Darwinian principle of infinite variation? Answer: (H + M)*4 and the physical principals of impenetrability and uncertainty. [ http://en.wikipedia.org/wiki/Lactate_dehydrogenase ]

Cheers, and only take 'biology' once daily before bed, and, with a grain of 'savor',

Fred Monson

Technical Director, CMIRT
West Chester University of Pennsylvania
Geology-Astronomy
West Chester, PA, 19383
http://cmirt.wcupa.edu

P.S. Is All histology artifact, or is there an exception outside of the undisturbed reality?

P.S. If all microscopic images originate in reciprocal space, what do we mean by the term, 'real image'?

_-----.---------------------
X-from: Elliott-at-arizona.edu [Elliott-at-arizona.edu]
Sent: Friday, May 11, 2012 9:02 PM
To: Monson, Frederick

Hi Fred

Of course there is not "10% of the molarity". That is clear to all.
The formula I suggested using is that if you take 10% of the value of
the molarity of formaldehyde, add it to 30% of the value of the
molarity of glut and add that to the molarity of the buffer, you will
get close to the effective molarity of your fixative.
For example, if you have a 4% solution of formaldehyde, you have a 40g/
L solution or approximately a 1.3M solution. 10% of THAT NUMBER gives
you about 0.13 effective.
It really is very simple, and not at all in need of being told, in
public, that % and molarity are different things.
The person I sent this to understood what I said. Also, several other
people who contacted me understood the formula. Even if you don't
agree with it (it has worked well for me for years), now at least you
understand it.

David

On Apr 12, 2012, at 3:30 PM, Fred Hayes wrote:

} There is no such thing as 10% of the molarity or 30% of the molarity.
} Molarity is expressed as moles/liter or moles of solute/liters of
} solvent. %
} is simply n parts of this vs 100 parts of that. % is concentration
} and has
} nothing to do with molarity. Two different ways of looking at
} solutions
} (solutes/solvent)

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The Sars International Centre for Marine Molecular Biology in Bergen,
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invertebrate Platynereis dumerilii.

Please follow http://sars.no/jobs/postdoc_hausen12Sars_04.php for more
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Closing date for written application is 22 June 2012.

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Email: SchottlerF-at-vision.wustl.edu
Name: Frank Schottler

Organization: Washington University, St. Louis, Ophthalmology

Title-Subject: Araldite embedding problem

Message: I have been using the same infiltration/embedding protocol for retinal eyecups using a modified Molenhauer Araldite 6005 formula for over 3 yrs without any problems until about 2 months ago. In the last couple of months the densest part of the tissue (ONL) acts as if it was incompletely penetrated or cured. I have replaced all chemicals as well as the dehdration/infiltration solvent (acetone) to no avail, and it does not seem that WPE has changed in the Epon 812 equivalent or the Araldite 6005.
Symptoms seen in 0.5 micron semithin sections from 3x3 mm blocks include, poor and faster overstaining with Toluidine blue, loss of detail in nuclei and variable and uneven staining of non-nuclear cytoplasmic regions. Increased stress fractures are observed in sequential 0.5 micron sections (not seen previously)and it is difficult if not impossible to get mirror-like block surface even after taking several ultra-thin sections(this is not a knife problem as tissue embedded 6 months ago cuts fine).
One suggestion has been to use the Molenhauer recipes posted on the vendor sites. However, the original protocol I inherited (which has worked for almost 3 yrs) differs from the Molenhauer recipes posted on the vendor sites in that the concentrations of 812-equivalent and 6005 are opposite to what are currently published (the 6005 being approximately twice that of 812). This concerns me, because of the potential issue of increased stress fracturing after using a higher concentration of 812, as thick sections (0.5-1 micron) of relatively large blocks (3x3 mm) are routinely needed.
The current protocol uses acetone as a dehydradting and infiltrating agent, and also uses an overnight infiltration with acetone/plastic (1:2) without the catalyst BDMA in the resin; the catalyst BDMA is added to the final 3 x 2hr changes prior to embedding on the subsequent day.
Thus, a related suggestion is to use ethanol/propylene oxide instead of acetone as a dehydradting and infiltrating agent. Obviously, including BDMA in all resin infiltrtion stages could also affect the final embedding quality.
I am at a loss to make any predictions, since it is unclear as to why the protocol stopped woking. ANY SUGGESTIONS?

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Email: SchottlerF-at-vision.wustl.edu
Name: Frank Schottler

Organization: Washington University, St. Louis, Ophthalmology

Title-Subject: Araldite embedding problem

Message: I have been using the same infiltration/embedding protocol for retinal eyecups using a modified Molenhauer Araldite 6005 formula for over 3 yrs without any problems until about 2 months ago. In the last couple of months the densest part of the tissue (ONL) acts as if it was incompletely penetrated or cured. I have replaced all chemicals as well as the dehdration/infiltration solvent (acetone) to no avail, and it does not seem that WPE has changed in the Epon 812 equivalent or the Araldite 6005.
Symptoms seen in 0.5 micron semithin sections from 3x3 mm blocks include, poor and faster overstaining with Toluidine blue, loss of detail in nuclei and variable and uneven staining of non-nuclear cytoplasmic regions. Increased stress fractures are observed in sequential 0.5 micron sections (not seen previously)and it is difficult if not impossible to get mirror-like block surface even after taking several ultra-thin sections(this is not a knife problem as tissue embedded 6 months ago cuts fine).
One suggestion has been to use the Molenhauer recipes posted on the vendor sites. However, the original protocol I inherited (which has worked for almost 3 yrs) differs from the Molenhauer recipes posted on the vendor sites in that the concentrations of 812-equivalent and 6005 are opposite to what are currently published (the 6005 being approximately twice that of 812). This concerns me, because of the potential issue of increased stress fracturing after using a higher concentration of 812, as thick sections (0.5-1 micron) of relatively large blocks (3x3 mm) are routinely needed.
The current protocol uses acetone as a dehydradting and infiltrating agent, and also uses an overnight infiltration with acetone/plastic (1:2) without the catalyst BDMA in the resin; the catalyst BDMA is added to the final 3 x 2hr changes prior to embedding on the subsequent day.
Thus, a related suggestion is to use ethanol/propylene oxide instead of acetone as a dehydradting and infiltrating agent. Obviously, including BDMA in all resin infiltrtion stages could also affect the final embedding quality.
I am at a loss to make any predictions, since it is unclear as to why the protocol stopped woking. ANY SUGGESTIONS?

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Hi

Just to keep it simple high voltage instability in an SEM would be best
displayed on a slow scan, the image moving in and out of focus. It is
extremely rare for this type of instability to appear due to the final lens
being unstable. I have only seen the final lens unstable once in 40 years
and that was a prototype!

Tell us more if your instrument does not follow the above.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
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To: protrain-at-emcourses.com

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Below is the result of your form, submitted on Friday, May 11, 2012
at 11:13:40 PM.

realname - Arazeshi
Email - {mailto:arazeshi-at-gmail.com} arazeshi-at-gmail.com
ORGANIZATION - Azad university
EDUCATION - Undergraduate College
LOCATION - Mashhad, Iran
SUBJECT_OF_QUESTION - SEM
QUESTION - What effects may be occur if we have high voltage
instability in an SEM? What are the diagnostic effects?

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Product Specialist - Microscopy

The Company:
Our client is a leading developer and producer of innovative high-tech
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The Opportunity:
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Salary:� Commensurate with�experience

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Primary Responsibilities:
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candidate will support trade shows and training courses, including
planning, communicating, attending and reporting.� Handling demonstration
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Education and Experience Required:
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preferred
- Business aptitude required
- Familiarity with biological imaging applications needed
- Fluorescence microscopy experience highly valued
- Good written and verbal communication skills are necessary

If you meet and/or exceed the experience criteria, please submit your
resume by emailing Christy Edwards (Sr. e-Recruitment Consultant with
Personify) at ce-at-personifysearch.com. We wish everyone the best of luck.
Unfortunately only qualified candidates will be considered.

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Dear colleagues,

I am writing to remind all interested researchers that the deadline for the
early-bird registration is in 5 days.

Event: Summer school on Micro- and Nano- structural characterization of
materials, focused on electron microscopy

Thanking you in advance,
Prof. E.K. Polychroniadis
Department of Physics, Aristotle University of Thessaloniki
Chairman of the Organizing Committee

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Here's a summary of the answers to the question I asked about SEM-EDS geometry.

Here's the question in abbreviated form: "...what take-off angle and working distance should he use?" I realize I should have referred to sample tilt instead. Take of angle is defined by working distance and "bore axis" (to borrow other terminology) of the mounted EDS detector.

What I was looking for was information on should you use the beam/sample/detector geometry recommended by the EDS manufacturer for your specific SEM or not. The only two parameters that I see as variable are sample tilt and working distance.

I want to thank everyone who took time to respond and send diagrams, information and references.

Here we go...

"...the proper distance would be where you get the maximum count rate. You would then need to determine the take-off angle and enter that with the EDS parameters."

"...instruments have character, no two from the same manufacturer are exactly the same, you always need to search for the "perfect" x-ray working distance.

How should you fix the problem? Well you need to find the WD that gives the highest x-ray signal as only in that position will the most trace element data be visible."

"...simple matter of determining the analytical working distance. I don't think a tilted sample should factor into that much.

Once there, I suppose you could play around with tilt and see where you get maximum counts. You would probably want to check the quant calibrations carefully to make sure they give decent results. I don't expect factory standards to give the right results. I am happily surprised when they are close. It would be better to collect their own.

I'd like to think that sample tilt can be factored into the quant routine. However, I doubt that tilt will really benefit them that much."

"Is this a trick question?" (No, I'm really curious about this. I was trained to have a flat sample at a specific working distance matching geometry predetermined by the SEM and the EDS manufacturer...FSK)

"I now realize that you were talking SEM, with variable geometry, I come from EPMAs, with fixed geometry, so I thought your question was "what takeoff angle and working distance should I enter into the analytical software?""

"Generally if it's angled down, the specimen is run level and the best WD is where the axis of the detector intersects the axis of the beam. Keeping the specimen horizontal (perpendicular to the beam) gives the smallest interaction volume for the x-rays. Tilting it gives more counts but generally from a larger part of the surface due to the beam running at an angle under the surface."

"I can see that tilting the sample might yield higher counts in this situation, but there is the drawback that you will be getting x-rays from a slightly larger area.

There is a second drawback to tilting the sample that I just remembered and that is that as you move around the sample you will need to keep moving the Z axis to maintain a constant WD.

Most systems will allow you to input the geometry data and they will then calculate the take-off angle and the solid angle for the quant corrections."

Again, Thank You! to everyone and if I'm misrepresenting your quote, tell me and I'll admit to it.

The majority seems to feel that due to each instruments little variations, the best approach is to find the best working distance that gives you the best/most x-ray counts, calculate the take-off angle and enter it into the software. Titled samples don't affect validly of the analysis, just decrease the spatial resolution.

Stay safe................
Frank

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Hi

We are considering moving up to a 200KV TEM. It has been a really long time since I have had to budget for something like this and I am hoping you might be willing to share some info with me to get started.

I am wondering if there is any current rule of thumb regarding what to expect to pay for a service contract for a fairly sophisticated instrument. And, if there might be any other 'hidden' charges like special all KV alignments that could turn up in addition to the cost of the contract.

Sorry to say I don't have much experience with anything over 120KV, just trying to get up to speed with current costs.

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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Colleagues, Friends,

Does anybody have practical experience using LMP agarose to immobilize
a pellet that won't stay together? Any advice on handling? Which type
do you prefer, VII or IX?
What concentration?
Finally - what is the temperature of agarose with which you mix your
sample?

I got these DNA nanoparticles that collaborators absolutely need
sectioned but can't produce in large amount. Tried to immobilize them
on a surface, but GA does not seem to work to secure them (there is
some protein-like polymer involved, but it must be not exposed). My
plan is now to get a maximally concentrated suspension and mix it with
a drop of gel-forming substance. Gelatine, my otherwise favorite,
solidifies too quick when in a small drop, and also shows quite a
presence under EM. Agarose is thinner, never liked how it handles so
don't have enough experience, but think it will mix easier with the
particle suspension, besides also being more clear. There is a concern
about temperature, though - I seem to remember, it took more heating
to liquify the agarose, compared with gelatin?..

Many thanks,
Vlad

________________________________________________
Vlad Speransky, Staff Scientist
Biomedical Engineering and Physical Science Shared Resource
National Institute of Biomedical Imaging and Bioengineering, NIH
13 South Dr, Rm. 3N17 MSC 5766
Bethesda, MD 20892
301 496-3989
vladislav_speransky-at-nih.gov
http://www.nibib.nih.gov/Research/Intramural/SharedResource/Speransky

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Robin

The Microscopy Society of America's Policy on Digital Imaging can be
found here

http://microscopy.org/resources/digital_imaging.cfm

I have reproduced it below.

The MSA position on digital image processing has been approved as follows:

"Ethical digital imaging requires that the original uncompressed image file be stored on archival
media (e.g., CD-R) without any image manipulation or processing operation. All parameters of the
production and acquisition of this file, as well as any subsequent processing steps, must be
documented and reported to ensure reproducibility.

Generally, acceptable (non-reportable) imaging operations include gamma correction, histogram
stretching, and brightness and contrast adjustments. All other operations (such as Unsharp-Masking,
Gaussian Blur, etc.) must be directly identified by the author as part of the experimental
methodology. However, for diffraction data or any other image data that is used for subsequent
quantification, all imaging operations must be reported."

This policy was formulated by the Digital Image Processing & Ethics Group of the MSA Education
Committee and was adopted as MSA policy at the Summer Council meeting August 2-3, 2003."

Cheers

Your Friendly Neighborhood SysOp

Nestor

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--
===========================================
Dr. Nestor J. Zaluzec
Argonne National Laboratory
Electron Microscopy Center
Materials Science/Bldg 212
Argonne, Illinois 60439 USA

Tel: 1-530-NES-TORZ (1-530-637-8679)
Fax: 1-630-252-4798

Email: Zaluzec-at-aaem.amc.anl.gov

iChat:Zaluzec-at-AIM
Skype: Zaluzec
TPM: http://tpm.amc.anl.gov
WWW: http://www.amc.anl.gov

Senior Scientist - Argonne National Laboratory
Fellow of the Microscopy Society of America
E.P. Wigner Fellow - Oak Ridge National Laboratory
Senior Fellow of the Computational Institute - University of Chicago
Past-President Microscopy Society of America

===========================================

The box said ...
"This program requires Win 95/98/NT/2K/XP/Vista or better..."
So I bought a Mac !

===========================================

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Hello Robin,
For imaging in biosciences, some years ago, an Editorial and a Feature on this topic were published in JCB (The Journal of Cell Biology):

Rossner, M. and Yamada, K. M. (2004) What’s in a picture? The temptation of image manipulation. J Cell Biol 166, 11-15.
doi: 10.1083/jcb.200406019

Rossner, M. ( 2008) A false sense of security. J Cell Biol, 183, 573-574.
doi: 10.1083/jcb.200810172

Best regards Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

On Friday 18 of May 2012 15:32:39 you wrote:
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Robin,

You might find this resource developed at UAB (funded by an Office of Research Integrity grant) to be of help: http://www.uab.edu/researchintegrityandimages/default.html

In addition to the MSA official policy that Dr. Zaluzec referred you to and the excellent articles by Michael Rossner cited by Oldrich, you might want to look at the Journal of Cell Biology's instructions to authors (particularly with regards to images) which have been a model for other journals (see: http://jcb.rupress.org/site/misc/ifora.xhtml). The Nature publishing group also is pretty specific (see: http://www.nature.com/authors/policies/image.html).

I have written some digital image ethics guidelines, which are available here: http://swehsc.pharmacy.arizona.edu/exppath/micro/digimage_ethics.php These guidelines were published as a full length article in 2010 (http://www.springerlink.com/content/00311qw26613m261/?p=499e7ec2a1174fad863e7b597298edd4&pi=0&MUD=MP)

Best regards,
Doug

^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^
Douglas W. Cromey, M.S. - Assistant Scientific Investigator
Dept. of Cellular & Molecular Medicine, University of Arizona
1501 N. Campbell Ave, Tucson, AZ 85724-5044 USA

office: AHSC 4212 email: Cromey-at-Arizona.edu
voice: 520-626-2824 fax: 520-626-2097

http://swehsc.pharmacy.arizona.edu/exppath/
Home of: "Microscopy and Imaging Resources on the WWW"

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Friday, May 18, 2012 6:31 AM
To: Cromey, Douglas W - (dcromey)

For a concise summary try:

http://www.igb.illinois.edu/sites/default/files/upload/core/PDF/Digital%20Im
aging%20Ethics.pdf

Not ethics per se, but also useful for accepted procedure:

ASTM E2765 - 11 Standard Practice for Use of Image Capture and Storage
Technology in Forensic Document Examination
ASTM E2339 - 11 Standard Practice for Digital Imaging and Communication in
Nondestructive Evaluation (DICONDE)

Tony
.........................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
Environmental, Health & Safety, Microscopy, & Forensic Engineering
pH2, LLC
5250 E US Highway 36, Suite 830
Avon, IN 46123
(317) 718-7020 Office
(317) 718-7038 Fax
(317) 409-3238 Cell
aahavics-at-pH2LLC.com
www.ph2LLC.com

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Sent: Friday, May 18, 2012 9:35 AM
To: ph2-at-sprynet.com

I also forgot one interesting sidebar:

Krueger, Forensic Examination of Questioned Scientific Images, Accountable
in Research, 9, 2, 105-125, 2002

This paper discusses the Office of Research Integrity's (ORI) experience
from its
oversight review of institutional scientific misconduct cases involving
questioned
images, provides illustrative examples of computer image processing
techniques
with an explanation of how the image analysis contributed to the final
misconduct
determination in specific cases, and reports a compilation of some generic
features of 35 ORI cases ( 1989- 200 1). The absolute and the relative
incidence of
cases with questioned images has increased over the last decade, as has the
use of
computers for presentation of the questioned image data. To date, ORI has
seen
no cases of de novo fabrication of images, and fortunately, in most cases
the
appearance of the questioned image has been quite secondary to the original
basis
for challenging its authenticity. The survey of cases indicates that good
laboratory
practices and, notably, attention to mentoring could have minimized the
scope
of the scientific misconduct when the act of falsification occurred.

Tony
.........................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
Environmental, Health & Safety, Microscopy, & Forensic Engineering
pH2, LLC
5250 E US Highway 36, Suite 830
Avon, IN 46123
(317) 718-7020 Office
(317) 718-7038 Fax
(317) 409-3238 Cell
aahavics-at-pH2LLC.com
www.ph2LLC.com

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com
[mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Friday, May 18, 2012 9:35 AM
To: ph2-at-sprynet.com

Around the end of March I posted that question about a PGT detector that was loosing calibration and generally not acting right. Time to (again) thank those who responded, and report the outcome.

A number of people counseled me to upgrade to a modern detector, but a couple (Thanks Mike!) suggested that Doug Connors, to whom Jim Nicolino sold his business when he retired, might be able to just bake it out and re-establish good vacuum. For future reference:

Douglas R Connors
AAT LLC.
TNAS Inc.
(608) 798-2005
(608) 7981675 fax
doug-at-tnas.net
www.tnas.net {http://www.tnas.net/}

I sent Doug the detector, he and his colleague re-evacuated it, tested it, found a couple of intermittent electrical "bad connections", fixed it, and sent it back - working like new.

I am delighted. The detector works like new, and cost was about 1/6 of what a new detector would have been, which pleases my frugal nature. Thanks Doug!

Now - if you are over 30 you can stop reading, but I think there may be some young people on the Listserv - the following is for them.

Everyone who said to replace / upgrade was right; I was cheap, stubborn, and wrong. Being cheap with your own money is wise; being cheap with your boss's money is nothing but silly. If you have a cat you know the motto: "It never hurts to ask for what you want." Worst they can say is "no" - so go for it!

In business, budget is a measure of status. Your boss may squeal like a pig being disemboweled when you suggest buying an expensive piece of equipment, but if you can justify it to him, and he can do the same, to his management - all that happens when he goes over budget is that next year he gets a bigger budget. This enhances his status. He may give you grief about it, but it advances his career. This principle may not be operative in academia, but it holds in the Corporate culture. Just sayin'

Have a great weekend!

Regards,
Andrew Werner
Chief Metallurgist, Perforating
Schlumberger Reservoir Completions
14910 Airline Road
Rosharon, TX 77583

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------------------------------------------------------------------------------------------------------------------------
NEWS! OWLS 2012 Genoa July 3-6 www.owls2012.org
Deadlines: Deadline for abstract submission prolonged to June 4th. Deadline for Early bird registration prolonged to June 8th.

Bursaries: "As regards financial assistance, OWLS organising committee, thanks to the generous support of ICO (www.ico-optics.org/) and EBSA (www.ebsa.org), has decided to provide 6 bursaries to assist the participation of students and young highly deserving scientists from developing areas. More details on the website soon."

Bursary application details: Two bursaries, US $ 750 each as provided by ICO, are mainly thought for extra EU participants. Four bursaries, 400 Euros each, will be directly provided by EBSA to EU participants. To apply, please, following your abstract submission, send a short CV and a short motivation to alberto.diaspro-at-iit.it, using as mandatory subject "owsl2012 bursary". Decision will be taken within June 15th according with prolongation of abstract submission deadline to June 4th. The winners will pay one half of the registration fee at the conference. In case registration has been already paid, the winner will be reimbursed at the conference of one half of the paid amount. We suggest to use the Early bird registration fee to save money for travel and lodging expenses.

Updated scientific program on the website www.owls2012.org
------------------------------------------------------------------------------------------------------------------------
Alby

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Dear colleagues,

We did change the LaB6 filament of our JEOL 3010 TEM last week and we did notice a problem with our spare wehnelt, during the cleaning process. The wehnelt cap's bore diameter was larger than I did remembered. We compared the diameter of it with the one from another TEM (JEOL 2100), and it was really big. The diameter was ~2.1mm compared to ~1.5mm. We did confirm that the wehnelt cap have the same part number for both microscope models. We do change filament once a year, and the microscope is 13-14 years old. I didn't see any reason to have the bore size enlarged that much.

Anyway, we did use this wehnelt as is to give a try. We couldn't adjust properly the gun tilt/shift properly with this wehnelt. So, we had to clean the wehnelt that was originally installed in the TEM and order a replacement cap.

After talking to the people here, I suspect that the person that was keeping the TEM before me was using a Dremel tool to clean and polish the wehnelt. More, I suspect he also did use some diamond polishing paste instead of the POL polishing compound to do the cleaning. Sure this will remove LaB6 deposit much faster than doing the traditional way by hand. But can someone over polish that much? Have this happened before to you?

Regards,

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

==============================Original Headers==============================
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6, 25 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
6, 25 -- Date: Mon, 21 May 2012 11:17:23 -0300
6, 25 -- Subject: TEM: Wehnelt cap bore size
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Hi Carlos

Yes I have seen a similar "problem" but in the case I investigated they had
chronic illumination astigmatism due to a very miss shaped cathode aperture.

Please understand that the cathode with a larger aperture may have been
appropriate for a tungsten hairpin system. A smaller cathode aperture may
have been used in order to attain the higher bias levels that LaB6 require.

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: carlos.inoki-at-lnls.br [mailto:carlos.inoki-at-lnls.br]
Sent: 21 May 2012 15:19
To: protrain-at-emcourses.com

Dear colleagues,

We did change the LaB6 filament of our JEOL 3010 TEM last week and we did
notice a problem with our spare wehnelt, during the cleaning process. The
wehnelt cap's bore diameter was larger than I did remembered. We compared
the diameter of it with the one from another TEM (JEOL 2100), and it was
really big. The diameter was ~2.1mm compared to ~1.5mm. We did confirm that
the wehnelt cap have the same part number for both microscope models. We do
change filament once a year, and the microscope is 13-14 years old. I didn't
see any reason to have the bore size enlarged that much.

Anyway, we did use this wehnelt as is to give a try. We couldn't adjust
properly the gun tilt/shift properly with this wehnelt. So, we had to clean
the wehnelt that was originally installed in the TEM and order a replacement
cap.

After talking to the people here, I suspect that the person that was
keeping the TEM before me was using a Dremel tool to clean and polish the
wehnelt. More, I suspect he also did use some diamond polishing paste
instead of the POL polishing compound to do the cleaning. Sure this will
remove LaB6 deposit much faster than doing the traditional way by hand. But
can someone over polish that much? Have this happened before to you?

Regards,

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

==============================Original Headers==============================
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"Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com} 6, 25 -- Date: Mon,
21 May 2012 11:17:23 -0300 6, 25 -- Subject: TEM: Wehnelt cap bore size 6,
25 -- Thread-Topic: Wehnelt cap bore size 6, 25 -- Thread-Index:
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} Hi Carlos
}
} Yes I have seen a similar "problem" but in the case I investigated they had
} chronic illumination astigmatism due to a very miss shaped cathode aperture.
}
} Please understand that the cathode with a larger aperture may have been
} appropriate for a tungsten hairpin system. A smaller cathode aperture may
} have been used in order to attain the higher bias levels that LaB6 require.
}
} Regards
}
} Steve

I think Steve is right. All of the LaB6 instruments we had came with two grid caps, one for W, one for LaB6. Sometimes it was hard to tell the difference.

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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Hi Carlos,
As one who has been using 1� diamond paste and a Dremel tool for most of the
past 30 years, I will say that, yes, it could be that the previous user was
over-enthusiastic, but I'm more inclined to think that your "spare" wehnelt
was for a tungsten cathode rather than a well worn wehnelt for LaB6,
especially if it is still symmetric.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: carlos.inoki-at-lnls.br [mailto:carlos.inoki-at-lnls.br]
Sent: Monday, May 21, 2012 10:23 AM
To: kenconverse-at-qualityimages.biz

Dear colleagues,

We did change the LaB6 filament of our JEOL 3010 TEM last week and we did
notice a problem with our spare wehnelt, during the cleaning process. The
wehnelt cap's bore diameter was larger than I did remembered. We compared
the diameter of it with the one from another TEM (JEOL 2100), and it was
really big. The diameter was ~2.1mm compared to ~1.5mm. We did confirm that
the wehnelt cap have the same part number for both microscope models. We do
change filament once a year, and the microscope is 13-14 years old. I didn't
see any reason to have the bore size enlarged that much.

Anyway, we did use this wehnelt as is to give a try. We couldn't adjust
properly the gun tilt/shift properly with this wehnelt. So, we had to clean
the wehnelt that was originally installed in the TEM and order a replacement
cap.

After talking to the people here, I suspect that the person that was
keeping the TEM before me was using a Dremel tool to clean and polish the
wehnelt. More, I suspect he also did use some diamond polishing paste
instead of the POL polishing compound to do the cleaning. Sure this will
remove LaB6 deposit much faster than doing the traditional way by hand. But
can someone over polish that much? Have this happened before to you?

Regards,

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

==============================Original Headers==============================
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Dear Colleagues,

Steve, that is a good point. At first I also thought it was a wehnelt for tungsten filament. But people told me that we never had such wehnelt for tungsten in our lab. I also talk to JEOL service engineer and he told me that the JEOL 3010 is to use only LaB6 filament.

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil
________________________________________
X-from: Steve Chapman [protrain-at-emcourses.com]
Sent: Monday, May 21, 2012 3:02 PM
To: Carlos Kazuo Inoki
Cc: Microscopical Soc of America

Dear colleagues,

We did change the LaB6 filament of our JEOL 3010 TEM last week and we did
notice a problem with our spare wehnelt, during the cleaning process. The
wehnelt cap's bore diameter was larger than I did remembered. We compared
the diameter of it with the one from another TEM (JEOL 2100), and it was
really big. The diameter was ~2.1mm compared to ~1.5mm. We did confirm that
the wehnelt cap have the same part number for both microscope models. We do
change filament once a year, and the microscope is 13-14 years old. I didn't
see any reason to have the bore size enlarged that much.

Anyway, we did use this wehnelt as is to give a try. We couldn't adjust
properly the gun tilt/shift properly with this wehnelt. So, we had to clean
the wehnelt that was originally installed in the TEM and order a replacement
cap.

After talking to the people here, I suspect that the person that was
keeping the TEM before me was using a Dremel tool to clean and polish the
wehnelt. More, I suspect he also did use some diamond polishing paste
instead of the POL polishing compound to do the cleaning. Sure this will
remove LaB6 deposit much faster than doing the traditional way by hand. But
can someone over polish that much? Have this happened before to you?

Regards,

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

==============================Original Headers==============================
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David:

7.5 liters / min

Richard E. Edelmann, Ph.D.
Center for Advanced Microscopy & Imaging, Director
Miami University
Oxford, OH 45056

Ph: 513.529.5712
www.cami.muohio.edu
________________________________________
X-from: zaluzec-at-microscopy.com [zaluzec-at-microscopy.com]
Sent: Tuesday, May 22, 2012 7:36 AM
To: Edelmann, Richard E. Dr.

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Name: David Lowry

Organization: Arizona State University

Title-Subject: water flow rates

Message: I am looking for the specified flow rates of chilled water
through the lens line and diff pump line on a JEOL JEM-1200EX TEM. Thank
you-

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Hello Listservers,
Does anyone have a clue on how important it is to separate HPF planchettes before placing the planchette in the AFS frozen cocktail. Some of our planchettes (hats) do separate (hexadecene coated), but some do not. At the end of the AFS run (5 days) all are separated. This question is related to my earlier one about a missing membrane AFS run, our very next AFS run did show membranes. We haven't pinned down the cause yet.

sincerely,
Michael Delannoy

==============================Original Headers==============================
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Hi Carlos

I visit many laboratories in a year and you would be surprised at what we
find when searching through the "spares". I am sure the information that
the service technician gives you is correct according to the JEOL
specification but as others have mentioned to cause such a remarkable change
in the cathode aperture has not been seen before.

My conclusion is that at some time a tungsten cathode has been used in the
laboratory, maybe an older instrument or delivered as a spare in error?

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: Carlos Kazuo Inoki [mailto:carlos.inoki-at-lnls.br]
Sent: 22 May 2012 13:04
To: protrain-at-emcourses.com
Cc: Microscopical Soc of America

Dear colleagues,

We did change the LaB6 filament of our JEOL 3010 TEM last week and we did
notice a problem with our spare wehnelt, during the cleaning process. The
wehnelt cap's bore diameter was larger than I did remembered. We compared
the diameter of it with the one from another TEM (JEOL 2100), and it was
really big. The diameter was ~2.1mm compared to ~1.5mm. We did confirm that
the wehnelt cap have the same part number for both microscope models. We do
change filament once a year, and the microscope is 13-14 years old. I didn't
see any reason to have the bore size enlarged that much.

Anyway, we did use this wehnelt as is to give a try. We couldn't adjust
properly the gun tilt/shift properly with this wehnelt. So, we had to clean
the wehnelt that was originally installed in the TEM and order a replacement
cap.

After talking to the people here, I suspect that the person that was
keeping the TEM before me was using a Dremel tool to clean and polish the
wehnelt. More, I suspect he also did use some diamond polishing paste
instead of the POL polishing compound to do the cleaning. Sure this will
remove LaB6 deposit much faster than doing the traditional way by hand. But
can someone over polish that much? Have this happened before to you?

Regards,

Carlos
__
Carlos Kazuo Inoki
Laborat�rio Nacional de Nanotecnologia
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

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Please remember! The original poster is most likely *not* on the microscopy mailing list! Please send you answers to the email address given with the question!
________________________________________

Below is the result of your form, submitted on Monday, May 21, 2012 at 08:05:24 AM.

realname - Georgianne Ciraolo
Email - Georgianne.Ciraolo-at-cchmc.org
ORGANIZATION - Cincinnati Chhildren's Hospital Medical Center
LOCATION - Cincinnati, Ohio 45229 USA
SUBJECT_OF_QUESTION - Plastic Embedding of mouse Bone
QUESTION - One our investigators would like to look at mouse bone. They want me to embed the samples in Osteo-Bed Bone embedding media. Does any one have any experience with this plastic. I have been told the the exothermic reaction might melt plastic embedding capsules. Is this true? Can I put the samples on ice during the infiltration steps?
Any advice would be appreciated.
Thanks to all
Georgianne Ciraolo
Georgianne.Ciraolo-at-cchmc.org

==============================Original Headers==============================
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Please see the following ad. Please note that this a state funded position.

-----------------------------------------------------------------------------------------------------------------------------------------------------------------
The Wyoming State Veterinary Laboratory, Department of Veterinary Sciences, University of Wyoming in Laramie is seeking an Electron Microscopist to support diagnostic and research activities at the university. The applicant will operate and maintain electron microscope (Hitachi H 7656 Transmission Electron Microscope), support equipment, and photographic equipment including an AMT digital camera. The incumbent will prepare diagnostic and research samples from a wide variety of biological tissues and fluids, using a variety of methods including, but not limited to, negative staining, thick and ultrathin sectioning.

In addition, the incumbent will perform and provide technical support for diagnostic pathology and virology service as well as provide research support for internal and external users. Instruct and train students, technical staff, and faculty and other users in EM lab procedures, protocols, methods, and techniques. Maintenance of accurate SOPs and quality control procedures is required to meet requirements for diagnostic laboratory certification.

Other duties will include general lab maintenance, monitoring inventory, order, receive, and distribute EM supplies; forecasting and planning laboratory needs, supervision of students and /or staff in the EM laboratory; setting priorities to meet EM project and lab work deadlines, designation work schedules and oversee accurate completion of assignments. This position will also oversee the operation of a central service laboratory, including training and scheduling of student hourly workers, ordering of supplies, and training of users.

Minimum Requirements: Requires a Bachelor�s degree or foreign equivalent in microbiology, biology, or a closely related field and relevant training or experience in microscopy. A Master�s degree and 2 or more years of experience involving TEM operation, including specimen preparation, digital imaging, and managing digital image archives is preferred.

The Wyoming State Veterinary Laboratory is a busy, full-service laboratory accredited by the American Association of Veterinary Laboratory Diagnosticians, providing testing for the diagnosis and surveillance of diseases in wild and domestic animals for clients in the State, intermountain region, and elsewhere. Research expertise in the Department of Veterinary Sciences includes infectious diseases occurring at the wildlife/domestic animal interface, toxicities in domestic animals and wildlife, and neurodegenerative diseases.

University of Wyoming is an EEO/AA employer. Background investigations are conducted on all prospective employees. Interested candidates can obtain more information on this position (Job ID #4923) and apply online at: https://jobs.uwyo.edu/.

Zhaojie Zhang, Ph. D.
Director, Jenkins Microscopy Facility
University of Wyoming
Laramie, WY 82071
PHONE: 307-766-3038
FAX: 307-766-5625

==============================Original Headers==============================
12, 30 -- From ZZhang-at-uwyo.edu Wed May 23 13:44:11 2012
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Unfortunately, one of the pneumatic vibration isolators on our scope blew out. is there a lister out there with a decommissioned 4000 who has such a part? We'd be willing to pay for the part and shipping.

Thanks!

Owen

Owen Mills
Michigan Tech University

==============================Original Headers==============================
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Dear All,
can anybody give me some advice with SEM preparation of insects (I am
working with honey-bees now and need to go through dehydration with
ethanol for final critical-point-drying), especially these topics:

- How to kill insects without getting these cramped legs? Use of
chloroform?
- Any tips how to bend and glue the legs of critical-point-dried
specimen to the stub?

Thanks,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

==============================Original Headers==============================
8, 20 -- From stefan.diller-at-t-online.de Fri May 25 05:28:37 2012
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Email: daisy.ridings-at-carolinashealthcare.org
Name: Daisy Ridings

Organization: Carolinas Medical Center

Title-Subject: TEM position available in Charlotte, NC

Message: Carolinas Medical Center in Charlotte, NC is seeking a full-time electron microscopy
technician. The position involves clinical and research work. Duties include operation and basic
maintenance of the transmission electron microscope, sample preparation, thick and thin sectioning,
staining, and digital imaging.

Qualifications: The candidate should have previous TEM experience. Electron Microscopy certification
preferred.Other required skills include the ability to troubleshoot problems with techniques and
equipment. Experience with digital imaging and Microsoft Office applications is highly desired.

Applicants must send your resume, history of your TEM experience, and the name and contact
information with 3 references to:

Helen E. Gruber, Ph.D.
Director, Orthopaedic Research Biology
Cannon Research Room 304
Carolinas Medical Center
PO Box 32861
Charlotte, N.C. 28232
704-355-5665; FAX: 704-355-5620
Helen.Gruber-at-carolinashealthcare.org

Carolinas Medical Center is an Academic Medical Center Teaching Hospital and the flagship facility
of Carolinas HealthCare System. Carolinas HealthCare System is an equal-opportunity not-for-profit,
self-supporting public organization offering a wide variety of health and human services to
residents of both North and South Carolina. Carolinas HealthCare System is the largest healthcare
system in the Carolinas, and the third largest public system in the nation.

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Email: gul417-at-mail.usask.ca
Name: Guosheng Liu

Organization: U of S

Title-Subject: Looking for Suppliers of parts for compound/dissecting microscopes

Message: Hello,

We got hundreds of compound/dissecting microscopes (mainly Olympus, from old CH models to newer
versions)for teaching but some of them are approaching at stage for service/repair. The most easily
broken parts, as I found, are stage holder spring and light bulb controller etc. It took me a lot of
time to contact original retailers but no positive feedback yet.

Does anybody know where it the best place to find these microscope-associated parts?

Btw, one of my clients is also looking for parts for this scope: see photo at
http://www.usask.ca/biology/scopes/MIcroscope%20Parts.pdf.

Any help will be greatly appreciated!

Guosheng

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All,

Can anyone suggest a laboratory (university preferred) that has in-situ
heating capability in TEM. I am also looking for in-situ
tensile/compression test.

Thanks
Vikram Bedekar

==============================Original Headers==============================
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-------- Original Message --------

I recently was given an old Sorvall MT-2 ultramicrotome in very good condition. The only problem with the microtome is that the 2 centimeter diameter "O" ring on the wheel between the hand crank wheel and drive wheel of the microtome had dry rot, and fell apart when I lubricated it. Now I need to replace the thing, and have no idea how to do so. Is there anyone on the listserver who has done so before and can guide me? I've been able to figure out how to replace all of the other belts on these microtomes in years past, but never needed to replace one of these before, so I don't know how to tackle the problem.

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046

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Electron Microscopy Core Facility - Deputy Director

Division of Intramural Research
National Heart, Lung, and Blood Institute
National Institutes of Health
Department of Health and Human Services

We seek an expert in the operation of scanning and transmission electron
microscopes and experience with electron tomography to serve as the Deputy
Core Director of the Electron Microscopy Core. The Core is in the Division
of Intramural Research (DIR), National Heart, Lung, and Blood Institute
(NHLBI), National Institutes of Health (NIH) in Bethesda, Maryland. The
successful applicant will interact with DIR Principal Investigators and
other scientific staff in providing advice, technical services and training
for all research requiring electron microscopy. In addition, the Deputy
Director will assist the EM Core Director in overseeing Core activities and
expanding the technical repertoire of the EM Core. More detailed
information about the NHLBI DIR may be found at http://dir.nhlbi.nih.gov/
{http://dir.nhlbi.nih.gov/}

Applicants must have a Ph.D. (or equivalent) in the biological sciences with
two or more years of postdoctoral research experience. Furthermore, the
successful candidate will have excellent interpersonal skills in working
with a large and diverse scientific community; familiarity with cell and
tissue ultrastructure; experience with the operation of scanning and
transmission electron microscopes; experience with electron tomography and
cryo-EM is desirable; hands-on experience with the techniques of sample
preparation for EM including two or more of the following: chemical and/or
cryofixation and embedding of cells and tissues, ultrathin sectioning of
embedded or rapidly frozen samples, immuno-gold labeling, negative staining,
rotary shadowing and metal replica preparation, freeze-etching/freeze
fracture, or critical point drying and metal coating for SEM. The successful
candidate will be expected to expand his/her technical repertoire to cover
most or all of these techniques during his/her tenure and will have the
opportunity to become familiar with a broad scope of biomedical research in
the NHLBI/DIR. Salary will be commensurate with qualifications and
experience.

Applicants should submit the following: cover letter highlighting key
qualifications, current curriculum vitae with complete bibliography, names
and contact information of three references, and a one-page summary of
applicant�s philosophy of core facility operation.

Applicants may be US citizens, resident aliens or non-resident aliens with
or eligible for a valid employment visa. Applications should be received by
July 15, 2012 but the advertisement will remain open until the position is
filled. PDF versions of documents sent by electronic mail are strongly
preferred. Materials should be sent to Dr. Mathew Daniels c/o Trina Gregory,
Administrative Officer, NHLBI, by email: gregoryp-at-nhlbi.gov.

HHS and NIH are Equal Opportunity Employers

Posted by Pat Connelly for Dr. Daniels

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E � Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov

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Hello,
I am informing myself about currently offered 120kV LaB6 TEMs and would appreciate if you would share your opinion on this machines with me.
1.) Would you recommend a particular model because of its known stability to act as a working horse in nanoparticle and biological thin section imaging? Having already one JEOL TEM in house, it would be an economic argument to stay with JEOL, in order to then share the already present cryo-transfer holders and equipment. I thus would compare all your recommendations with the JEOL candidate, a JEM-1400 HC + cryo shields. But we also have good reasons to change to a different brand now, especially because of our very bad experience with the long down times of our JEOL equipment due to the horribly long delays in spare part deliveries and subsequent delays in receiving repairs fulfilled in acceptable times. So, how does a FEI Spirit (TWIN) in your opinion compares to a JEM-1400, and would ZEISS or HITACHI have something to offer in this price range?
2.) Would you have a recommendation for which bottom mounted CCD camera to go for?
I at the moment thick about a TVIPS TemCam-F216 (2k x 2k, 32 x 32 mm2), and this would be for sure the most expensive device I could go for. But I am not aware about more economic and still somehow competitive cameras. I am afraid that our financial plan will finally not allow me to go for the TVIPS camera, and therefore I also need to get known about the good alternatives.

What are we going to measure with our new 120 kV LaB6 TEM?
Imaging biofunctionalized nanomaterials, I search for _very high contrast_ (organic/biological materials, ultrathin resin sections, cryo-EM) and still _very high resolution_ (imaging size distributions of gold nanoparticles { 2 nm diameter). I know, that contradicts, and therefore a good compromise is what I need, and therefore I am asking for your recommendations. There will no EDX or EELS, and no STEM being installed on the new 120 kV LaB6 machine. A multi-grid retainer sample holder shall be used for achieving high throughput in the nanoparticle size distribution analysis, and a cryo-transfer sample holder for cryo-EM at low dose illuminations will be run on that machine. Tomography will not be done. Acquiring images for single particle reconstructions might become a topic in the future. We already have a JEOL JEM-2100F 200kV FEG TEM with ultra high resolution polepiece and STEM and EDX for our high end works. We now need to amplify our laboratory by a second TEM for outsourcing the most numerous but not-so-high-end routine work from that 200kV machine to a very economic new 120 kV machine.

Thanks for your recommendations, and best greetings from SPAIN,
Marco M�ller

----------------------------------------------------------------
Marco M�ller
Platform Manager - Electron Microscopy

CIC biomaGUNE
Parque Tecnol�gico de San Sebasti�n
Paseo Miram�n, 182. Ed. Empresarial C
20009 San Sebasti�n (Guip�zcoa)
SPAIN

Tel:� +34 943 00 53 25
mmoller-at-cicbiomagune.es

This e-mail is from CIC biomaGUNE. The e-mail and any files transmitted with it are confidential and intended solely for the use of the individual or entity to whom they are addressed. Any unauthorised dissemination or copying of this e-mail or its attachments, and any use or disclosure of any information contained in them, is strictly prohibited and may be illegal. If you have received this e-mail in error, please notify or telephone + 34 943 00 53 00 and delete it from your system.

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Ed,

You can get at this wheel by removing the hand-crank, then the case,
and the crank wheel inside. Be sure to run the small wheel to one end
of its range first!
I'd get more detailed, but I have to first haul my parts 'tome out of
the cabinet ...
You might also want to try the open cut list:
open_cut_project-at-googlegroups.com

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For those of us who have orphaned ultratomes (and who doesn't?).

Phil

} I recently was given an old Sorvall MT-2 ultramicrotome in very good
} condition. The only problem with the microtome is that the 2
} centimeter diameter "O" ring on the wheel between the hand crank
} wheel and drive wheel of the microtome had dry rot, and fell apart
} when I lubricated it. Now I need to replace the thing, and have no
} idea how to do so. Is there anyone on the listserver who has done so
} before and can guide me? I've been able to figure out how to replace
} all of the other belts on these microtomes in years past, but never
} needed to replace one of these before, so I don't know how to tackle
} the problem.
}
} Ed
}
}
} Edward Haller, Lab Manager
} University of South Florida
} Integrative Biology Department
} Electron Microscopy Core
} SCA 110
} 4202 East Fowler Avenue
} Tampa, FL 33620
} 813-974-2676
} ehaller-at-usf.edu
} Office: ISA 1046

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Dear,
within the LANIR project (webpage under construction: http://www2.ul.ie/web/Research/Lanir) I am offering a post doc position, immediately available, for two years at http://www.iit.it/en/research/departments/nanophysics.html. The call will be released soon on the IIT webpage. I need the candidate has expertise in optical microscopy image formation, NIR/IR laser systems, NIR/IR spectroscopy.
Please, feel free to write for information to alberto.diaspro-at-iit.it (copy to manuela.salvatori-at-iit.it), subject: "LANIR-IIT Diaspro Lab". A "Job corner" for interviews will be available during www.owls2012.org at the IIT/LANIR stand.
All the best
Alby

Note: Within the LANIR project we outline a concept of Infra Red Nanoscopy (IRN). It is expected that IRN has the potential for improving the lateral resolution of IR micro-spectroscopy from the diffraction-limited current state-of-the-art down to the nanoscale. Main applications are expected in Medicine, Biology and Materials science.
The LANIR research leading to these results has received funding from the European Community's Seventh Framework Programme (FP7/20012-2015) under grant agreement n�280804. This communication reflects the views only of the author, and the Commission cannot be held responsible for any use which may be made of the information contained therein.

ISTITUTO ITALIANO
DI TECNOLOGIA

Prof. Alberto Diaspro
Director
Department of Nanophysics
Istituto Italiano di Tecnologia
Via Morego, 30 16163 Genova
Tel: +39-010.71.781.503
Fax +39-010-72.03.21
Mobile +39-3666719968
www.iit.it
alberto.diaspro-at-iit.it

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Hello Listers,
We will need to purchase a new glass knife breaker. I am looking for any comments on breakers which are currently on the market. What is available, what are your experiences with them.
The knives we will be making for the ultramicrotome are primarily for 0.45 micron sections and for facing blocks before ultrathin sections. We use 6.4 mm glass.

Thank you,
Rosey

Rosey van Driel
Manager, Transmission Electron Microscopy
Institute for Frontier Materials
Geelong Technology Precinct
Deakin University,�
75 Pigdons Rd
Waurn Ponds, Vic 3217,� Australia

Telephone: +61 3 5227 3117
Facsimile: 03 5227 1103
Email:rosey.vandriel-at-deakin.edu.au
Website:http://www.deakin.edu.au
Deakin University CRICOS Provider Code 00113B

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Hi Rosey,

I make our knives with a Leica EM KMR3 knife maker, with 6.4
mm glass. I use glass knives primarily for block facing, but
the knife edges are excellent for thin sectioning as well. I
have used Reichert-Jung and RMC knife makers (older models)
in the past and much prefer the KMR3 because its design is
so much more user friendly, and I get consistently good
knives with it. I would highly recommend it.

Cheers,

Gigi

�
Gigi Kemalyan
Nogales Lab Research Technician
Howard Hughes Medical Institute
University of California, Berkeley
742 Stanley Hall
Berkeley, CA 94720-3220
Ph 510-666-3335
Email (work): gigik-at-berkeley.edu
Email (alt.): singinggardenersx2-at-live.com
Website: http://cryoem.berkeley.edu/

�

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Hello Listers,

We will need to purchase a new glass knife breaker. I am looking for any
comments on breakers which are currently on the market. What is available, what
are your experiences with them.

The knives we will be making for the ultramicrotome are primarily for 0.45
micron sections and for facing blocks before ultrathin sections. We use 6.4 mm
glass.

Thank you,
Rosey

�
Rosey van Driel
Manager, Transmission Electron Microscopy
Institute for Frontier Materials
Geelong Technology Precinct
Deakin University,�
75 Pigdons Rd
Waurn Ponds, Vic 3217,� Australia
�
Telephone: +61 3 5227 3117
Facsimile: 03 5227 1103
Email:rosey.vandriel-at-deakin.edu.au
Website:http://www.deakin.edu.au
Deakin University CRICOS Provider Code 00113B

-------
No trees were harmed by sending this message; however, a large number of electrons were greatly inconvenienced.

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Dear All,

About 45 minutes into the film a TEM comes into shot sitting in one of the
corridors of the ship Prometheus (stability of spacecraft in 2093 must be
pretty good). I reckon it's a JEOL JEM1400 with a paint job. The really
interesting thing about this machine is that it can read alien DNA using
what looks to be a little green laser mounted on the beam-stop! Very cool!
Not only that, it can send the DNA profiles direct to their mobile phones
("index this" app?).

The one sad thing is that their scope still has manual aperture drives
(reliability issues? Can't get a service engineer several lights years
out?) and it still has a viewing chamber. I guess some technology is too
good to leave behind. Nice to see TEM still representing the kick-ass end
of technology far into the future.

Best, Jon

P.S. I didn't spot any mention of JEOL in the end credits. Any ideas?

P.P.S. My other half gave audible groan when I got excited to see a TEM in
the film. :)

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X-from: Yang, Ann-Fook {Ann-Fook.Yang-at-AGR.GC.CA}

Hi Stephen,

The legs become flexible after critical point drying. You may arrange them anyway you want before
gold coating.

Ann-Fook Yang,
Microscopy Unit | Unite de Microscopie
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
Edifice K.W. Neatby Building,
960 Carling Av
Ottawa,Ontario
K1A 0C6
ann-fook.yang-at-agr.gc.ca
Telephone | T�l�phone: 613-759-1638
Facsimile | T�l�copieur: 613-759-1701
Teletypewriter | T�l�imprimeur 613-773-2600
Government of Canada | Gouvernement du Canada

-----Original Message-----
X-from: stefan.diller-at-t-online.de [mailto:stefan.diller-at-t-online.de]
Sent: Friday, May 25, 2012 6:38 AM
To: Yang, Ann-Fook

Dear All,
can anybody give me some advice with SEM preparation of insects (I am working with honey-bees now
and need to go through dehydration with ethanol for final critical-point-drying), especially these
topics:

- How to kill insects without getting these cramped legs? Use of chloroform?
- Any tips how to bend and glue the legs of critical-point-dried specimen to the stub?

Thanks,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
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Name: David Reuschle

Organization: Dow Chemical

Title-Subject: Razor Blades

Message: I am considering some alternatives to razor blades for trimming polymer samples for TEM,
SEM, and AFM (block face, etc.). Dexterity is important, of course. Any suggestions or comments
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Email: amy.albin-at-utoledo.edu
Name: Amy Albin

Organization: University of Toledo

Title-Subject: I need another manual

Message: Everyone was so helpful the last time, I was wondering if I could see if anyone could help
me with my next manual quest. I am looking for a manual for a Dupont Instuments Sorvall JB-4
microtome... preferably in pdf form if possible. Thank you for your time and energy.

-Amy Albin
University of Toledo Electron Microscopy Lab

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Rough trimming before sectioning? Or ... ?
Rough trimming can be done with a glass knife on an ultratome or a
JB-4 microtome, or something like a LKB Pyrimatome.
If the glass transition temperature is high enough, you can also try
LN2 cryofracture. This produces a better face for SEM and probably
AFM, but won't help much for TEM.

Phil

} Email: dreuschle-at-dow.com
} Name: David Reuschle
}
} Organization: Dow Chemical
}
} Title-Subject: Razor Blades
}
} Message: I am considering some alternatives to razor blades for
} trimming polymer samples for TEM,
} SEM, and AFM (block face, etc.). Dexterity is important, of course.
} Any suggestions or comments
} are welcome!

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Hi listers,

I've been preparing TEM cross-sections using a tripod polishing
method--once the samples are fully thinned I'd like to glue down an
aperture grid for support. Currently we've been using some
store-bought five minute epoxies, but I'm concerned about sample
contamination and possible outgassing in our microscope.

Are there any epoxies that you would recommend for such an application?

Thanks for your suggestions.

--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering
Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

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Ophthalmology scalpels, both disposable and re-usable, are incredibly
convenient for small-scale work :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

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}
} Title-Subject: Razor Blades
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***************************************************************************************
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} realname - Tracey Pepper
} Email - tpepperisu-at-gmail.com
} ORGANIZATION - Iowa State University
} EDUCATION - Graduate College
} SUBJECT_OF_QUESTION - arabidopsis root fixation
} QUESTION - Hello all!
}
} I'm struggling with a project in the lab. We are looking at arabidopsis
} roots, 7 day old seedlings, about 1-1.5 cm from the tip. Cutting
} longitudinal sections for LM and TEM evaluation. My issue is that I cannot
} stop the plasmolysis (plasma membranes distending randomly from the cell
} wall....not in all cells, but still there in some)! I've tried many
} variations of buffers (cacodylate 0.025, 0.05, 0.085, 0.1 M) with
} variations of para/glut combinations to no avail. At this point, the
} 0.025M cacodylate buffer shows the least amount of plasmolysis combined
} with 1% para+1% glut fix. (pH 7.2) These are for general morphology, so we
} are osmicating in matching buffer at 1% for 0.5hr and dehydrating, thru
} acetone, into Spurr's. Any suggestions? The mutation is supposed to cause
} massive membrane disruptions, so seeing disruption in the WT is not a good
} place to start!
}
} Thanks in advance!!
}
} Tracey Pepper
} Microscopy and NanoImaging Facility
} Iowa State University
} Ames, IA

--

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Email: jinguo.wang-at-utdallas.edu
Name: Jinguo Wang

Organization: UT Dallas

Title-Subject: opinion on the Remi Group as an equipment service contract provider

Message: Dear All,

My institution suggested me to consider the Remi group for servicing our equipments. If anyone have
experience with the Remi group, I would appreciate to hear from you about the good and the bad about
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Jinguo Wang

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I use Epo-Tek 353ND from Epoxy Technology. It is equivalent to Gatan's G1
epoxy.

If you don't use tabbed grids and hold the grids with tweezers when you
lower the grid onto the sample, there is another way with standard grids.
Cut a Post-ItR note into a point with the sticky end at the point. Cut the
tip so that it is flat across. Put a bend on the Post-it note so that it
will be parallel to the surface when you lower into to press it onto the
grid. Paint your grid, blot it, and then lower the grid onto your sample by
simply holding the Post-It note. Works like a charm.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: spurgeon-at-drexel.edu [mailto:spurgeon-at-drexel.edu]
Sent: Friday, June 08, 2012 6:22 AM
To: s.walck-at-cox.net

Hi listers,

I've been preparing TEM cross-sections using a tripod polishing method--once
the samples are fully thinned I'd like to glue down an aperture grid for
support. Currently we've been using some store-bought five minute epoxies,
but I'm concerned about sample contamination and possible outgassing in our
microscope.

Are there any epoxies that you would recommend for such an application?

Thanks for your suggestions.

--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

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Hi Jon,

There are already projects running long before 2093 to sequence DNA with�TEM and AFM.
The people at JEOL must find it comforting to see that they will be chosen to install TEMs on the human interstellar flotilla in the future and my other half would be eager�to know if�aliens share the same DNA code as Earth species.

Stephane

----- Original Message -----
X-from: "jsb43-at-cam.ac.uk" {jsb43-at-cam.ac.uk}
To: nizets2-at-yahoo.com
Cc:
Sent: Tuesday, June 5, 2012 10:56 PM

Dear All,

About 45 minutes into the film a TEM comes into shot sitting in one of the
corridors of the ship Prometheus (stability of spacecraft in 2093 must be
pretty good). I reckon it's a JEOL JEM1400 with a paint job. The really
interesting thing about this machine is that it can read alien DNA using
what looks to be a little green laser mounted on the beam-stop! Very cool!
Not only that, it can send the DNA profiles direct to their mobile phones
("index this" app?).

The one sad thing is that their scope still has manual aperture drives
(reliability issues? Can't get a service engineer several lights years
out?) and it still has a viewing chamber. I guess some technology is too
good to leave behind. Nice to see TEM still representing the kick-ass end
of technology far into the future.

Best, Jon

P.S. I didn't spot any mention of JEOL in the end credits. Any ideas?

P.P.S. My other half gave audible groan when I got excited to see a TEM in
the film. :)

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Dear Colleagues,

I have done mainly conventional TEM an very little HRTEM, and limited to SI and III-V semiconductors. That's why I am looking for your advise. I do have a colleague doing HRTEM of nano particles. He is doing the image simulation using a commercial package to match the model of the nano crystal with his observations. To do the simulation several parameters are necessary. The question is how these parameters affect the simulation? I know that these parameters needs to me measured for each TEM, except maybe the Cs that is provided with the microscope (I suppose it is very close if I measure it). Now, this colleague of mine is using parameters provided with the simulator. And these parameters are not exactly for the microscope he is using. I insisted that the microscope has to be characterized to get the correct parameters, but he argues that he tried changing the simulation parameters and he doen't see much change in the results.

What do you think? How much you need to know about your instrument to be confident about a HRTEM image simulation?

Thanks,

Carlos

__
Carlos Kazuo Inoki
Brazilian Nanotechnology National Laboratory
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

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Email: michael.cammer-at-med.nyu.edu
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Organization: Skirball Institute of Biomolecular Medicine

Title-Subject: 50nm resolution of micropillars in live cell light microscopy

Message: Does anybody have experience with imaging the forces of cells as measured by the movements
of synthetic micropillars as a cell substrate? For instance, O du Roure "Force mapping in
epithelial cell migration" PNAS 102(7) reports "The resolution of the displacements is of the order
of 50 nm..." This is measured by imaging cells by brightfield, phase contrast, or Nomarski and the
tops of the micropillars the cells sit on form bright spots. The underlying spots move as the cell
pulls on the substrate. I was wondering how this can be calculated from images obtained with
microscope objectives of N.A. from 0.80 to 1.40. For instance, there seem to be MatLab scripts
floating around that do this accurately (but we'd rather use ImageJ). Any help would be appreciated
as we would like to use this method.
Thank you in advance.
Regards,
Michael

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Title-Subject: Best B&W printer for micrographs

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Hi Michael,

I didn't know the work of this group and it is quite interesting.
However I have the feeling that the critical part of the work, which is to precisely localize the top of the micropillars in order to determine the forces, is not described with enough details. For example I think that the resolution of the camera should be given because it gives an idea of the size of the pixel.
Also I regret that the error of measurement of the micropillar position is not accurately calculated and accurately described.
Since the authors calculate the position of the micropillars through extrapolation of the gaussian curve, there is an error associated with this determination and I see nowhere that this error is discussed or the method of calculation presented.�The pillars bend along their length, with their base mainly staying in place and only the top moving. An interesting question would be to study the contribution of the base vs. top of the pillar in the formation of the so-called "bright spots" which are actually analyzed.

I also found an interesting article discussing this point :
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2881340/

I want to say that I am not a specialist of this method and the best person to discuss it would be its inventor so perhaps I missed something but I think it is an interesting subject of discussion.

Best regards,
Stephane

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Email: michael.cammer-at-med.nyu.edu
Name: Michael Cammer

Organization: Skirball Institute of Biomolecular Medicine

Title-Subject: 50nm resolution of micropillars in live cell light microscopy

Message: Does anybody have experience with imaging the forces of cells as measured by the movements
of synthetic micropillars as a cell substrate?� For instance, O du Roure "Force mapping in
epithelial cell migration" PNAS 102(7) reports "The resolution of the displacements is of the order
of 50 nm..." This is measured by imaging cells by brightfield, phase contrast, or Nomarski and the
tops of the micropillars the cells sit on form bright spots.� The underlying spots move as the cell
pulls on the substrate.� I was wondering how this can be calculated from images obtained with
microscope objectives of� N.A. from 0.80 to 1.40.� For instance, there seem to be MatLab scripts
floating around that do this accurately (but we'd rather use ImageJ).� Any help would be appreciated
as we would like to use this method.
Thank you in advance.
Regards,
Michael

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I have a size fraction of {75um rock particles that I need to further
separate into } 45um, } 15um & {15um. I've tried wet sieving with precision
fabric, but because the 15um sieve has only 10% open space it took much
longer than anticipated, and still did not pass more than 40-50% of that
size. I'm looking into jet sieving, but thought I'd ask this group about
their experience with wet sieving, jet sieving, and other possible
methods(?)

TIA :o)
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
Information: http://www.mun.ca/research/ocp/creait/maf/
Scheduling: http://www.mun.ca/research/ocp/creait/maf/calendar.php
(709) 864-6799 (Ofc)
(709) 864-6790 (Lab)
cogito ergo ZzoooomM

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Hi Michael!

I don't know jet sieving but you are clearly in a delicate zone (between 100 and 10�m) :-)
Particules bigger than 10�m usually sediment fast in water (of course it depends on the density), so to select particles under this size�you just suspend the powder in water and leave the suspension for 5 min on the bench and then pour the supernatant.�Repeat the process 3x and�the results should be quite good.
For even smaller particles (sub-micro) centrifugation works well.
I suppose it may be possible to use higher density liquids (than water) in order to select bigger particles but I never experienced that.
Best regards,

Stephane

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Sent: Tuesday, June 12, 2012 6:15 PM

I have a size fraction of {75um rock particles that I need to further
separate into } 45um, } 15um & {15um.� I've tried wet sieving with precision
fabric, but because the 15um sieve has only 10% open space it took much
longer than anticipated, and still did not pass more than 40-50% of that
size.� I'm looking into jet sieving, but thought I'd ask this group about
their experience with wet sieving, jet sieving, and other possible
methods(?)

TIA� :o)
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
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Scheduling:�� http://www.mun.ca/research/ocp/creait/maf/calendar.php
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�� (709) 864-6790 (Lab)
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Hi all

It seems that we'll have to re-pump our EDS detector. I must now ask our
workshop to make the T tube with the tool to open the plug on the
detector and the pumping output.
I can good imagine how it look like, but if someone would have a squetch
to send me, it would be nice and save me some time, in particular as the
critical dimensions are probably in inches and not metric.

Thanks in advance

Jacques

--

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Mat�riaux de Strasbourg
D�partement Surfaces et Interfaces
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail : Jacques.Faerber-at-ipcms.unistra.fr

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Email: amit.welcomes.u-at-gmail.com
Name: Amit

Organization: Indian institute of science

Title-Subject: electron microscopy of magnetic samples

Message: Good evening,
We work on jeol jem 2100f. I was just browsing site of jeol and found out that they have mentioned
for JEM-2100 Lab6 that it has an objective mini lens hence lorentz microsopy is a default feature or
we can do magnetic samples in it.
3rd line in 4th paragraph:
http://www.jeol.com/products/electronoptics/transmissionelectronmicroscopestem/200kv/jem2100lab6/tabid/123/default.aspx

but same is not said about 2100f. as both have same lens configuration (other than one extra
condenser lens in lab6 system) shall i presume its true for 2100f also?
can anyone tell what is the gap between objective pole piece in HR objective pole piece in
JEM-2100F? can we image magnetic nao particles normally in it?(i e without switching off the
objective lens)

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Can someone remind me how this is done? I can't find the instructions online and I have misplaced the piece of paper that comes with the journal.

Thanks,
Doug

^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^
Douglas W. Cromey, M.S. - Assistant Scientific Investigator
Dept. of Cellular & Molecular Medicine, University of Arizona
1501 N. Campbell Ave, Tucson, AZ 85724-5044 USA

office: AHSC 4212 email: Cromey-at-Arizona.edu
voice: 520-626-2824 fax: 520-626-2097

http://swehsc.pharmacy.arizona.edu/exppath/
Home of: "Microscopy and Imaging Resources on the WWW"

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Amit-

I am very interested to know what you find out. We have a JEOL 2100 and I have just recently been looking for an explanation of the reference to Lorentz microscopy mentioned at the link you provided. On the features/capabilities document it says "Field Free Imaging - Lorentz Microscopy - Foucault/Lorentz - Standard". Also, "Lorentz mode standard: Foucault imaging from 100X to 40kX (inexpensive option)". In another place: "Low Mag optics/Lorentz Microscopy - Standard - simple upgrade to full Foucault imaging up to 40kX". So, they may be referring to a change of pole piece, which I don't really want to pursue, if it means lower resolution. Otherwise, I'm guessing they just mean to work in low-mag mode, (with the OL off), but that doesn't seem adequate for analyzing nanostructures. But if there is some software that would make it easier to work with magnetic samples, I would be very interested. I'm hoping to get some feedback from JEOL soon.

-Phil
------------------------------------------
Phil Ahrenkiel, Assistant Professor
Nanoscience and Nanoengineering Ph.D. Program
South Dakota School of Mines and Technology
501 E. Saint Joseph St.
Rapid City, SD 57701 U.S.A.
Office: EP 221
Phone: 605-394-5238, Fax: 605-394-2365
Email: Phil.Ahrenkiel-at-sdsmt.edu

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Email: amit.welcomes.u-at-gmail.com
Name: Amit

Organization: Indian institute of science

Title-Subject: electron microscopy of magnetic samples

Message: Good evening,
We work on jeol jem 2100f. I was just browsing site of jeol and found out that they have mentioned for JEM-2100 Lab6 that it has an objective mini lens hence lorentz microsopy is a default feature or we can do magnetic samples in it.
3rd line in 4th paragraph:
http://www.jeol.com/products/electronoptics/transmissionelectronmicroscopestem/200kv/jem2100lab6/tabid/123/default.aspx

but same is not said about 2100f. as both have same lens configuration (other than one extra condenser lens in lab6 system) shall i presume its true for 2100f also?
can anyone tell what is the gap between objective pole piece in HR objective pole piece in JEM-2100F? can we image magnetic nao particles normally in it?(i e without switching off the objective lens)

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On Jun 13, 2012, at 3:37 AM, jacques.faerber-at-ipcms.u-strasbg.fr wrote:

} Hi all
}
} It seems that we'll have to re-pump our EDS detector. I must now ask
} our
} workshop to make the T tube with the tool to open the plug on the
} detector and the pumping output.
} I can good imagine how it look like, but if someone would have a
} squetch
} to send me, it would be nice and save me some time, in particular as
} the
} critical dimensions are probably in inches and not metric.
}
} Thanks in advance
}
} Jacques
}
} --
}
} J. Faerber
} IPCMS-DSI
} Institut de Physique et Chimie des Mat�riaux de Strasbourg
} D�partement Surfaces et Interfaces
} 23, rue de Loess ; BP43
} 67034 Strasbourg CEDEX 2
} France
}
} Tel 00 33(0)3 88 10 71 01
} Fax 00 33(0)3 88 10 72 48

Dear Jacques,
When we had this issue, we replaced the back panel of the detector
with one that had a standard vacuum valve attached, then we could just
attach a line to pump out the detector. I do not know whether the
housing on your detector is capable of this modification, but it
solved our problem completely.
Yours,
Bill

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Thanks Stephane �

Others had suggested sediment rates as well. However, there exists
relatively large density differences between the minerals in these
powdered samples, the most important of which are quartz and
hematite/magnetite, and carbonates between. Subsequent analysis of the
modes would depend on there being absolutely no partitioning of minerals
relative to size.

Still investigating :o)

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Genuinely, Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
cogito ergo ZzoooomM

On 12-06-13 5:28 AM, "Stephane Nizet" {nizets2-at-yahoo.com} wrote:

} Hi Michael!
}
} I don't know jet sieving but you are clearly in a delicate zone (between
} 100 and 10�m) :-)
} Particules bigger than 10�m usually sediment fast in water (of course it
} depends on the density), so to select particles under this size you just
} suspend the powder in water and leave the suspension for 5 min on the
} bench and then pour the supernatant. Repeat the process 3x and the
} results should be quite good.
} For even smaller particles (sub-micro) centrifugation works well.
} I suppose it may be possible to use higher density liquids (than water)
} in order to select bigger particles but I never experienced that.
} Best regards,
}
} Stephane
}
}
}
} ----- Original Message -----
} From: "mshaffer-at-mun.ca" {mshaffer-at-mun.ca}
} To: nizets2-at-yahoo.com
} Cc:
} Sent: Tuesday, June 12, 2012 6:15 PM
} Subject: [Microscopy] separating the smallest of particles
}
}
}
}
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Title-Subject: NIS Elements Viewer installation help?

Message: Nikon uses InstaShield for packaging software at
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Hi Steven,

We use a small amount of Gatan G1 or M Bond 610 for gluing our tripod polished XTEM samples onto Molybdenum or Copper slot type grids.

Hope that is useful, if you need any assistance with this technique - please let me know.

Thanks,

Chris

Best Regards,
Christopher J. Pawley
Ph.D Student
Materials and Physics Research Centre, Room 111/114, Maxwell Building,
University of Salford, The Crescent, Salford, M5 4WT

________________________________________
X-from: S.Walck-at-cox.net [S.Walck-at-cox.net]
Sent: 10 June 2012 04:59
To: Pawley, Christopher (PG)

I use Epo-Tek 353ND from Epoxy Technology. It is equivalent to Gatan's G1
epoxy.

If you don't use tabbed grids and hold the grids with tweezers when you
lower the grid onto the sample, there is another way with standard grids.
Cut a Post-ItR note into a point with the sticky end at the point. Cut the
tip so that it is flat across. Put a bend on the Post-it note so that it
will be parallel to the surface when you lower into to press it onto the
grid. Paint your grid, blot it, and then lower the grid onto your sample by
simply holding the Post-It note. Works like a charm.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: spurgeon-at-drexel.edu [mailto:spurgeon-at-drexel.edu]
Sent: Friday, June 08, 2012 6:22 AM
To: s.walck-at-cox.net

Hi listers,

I've been preparing TEM cross-sections using a tripod polishing method--once
the samples are fully thinned I'd like to glue down an aperture grid for
support. Currently we've been using some store-bought five minute epoxies,
but I'm concerned about sample contamination and possible outgassing in our
microscope.

Are there any epoxies that you would recommend for such an application?

Thanks for your suggestions.

--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

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Hi Garry

I am relatively new to section/slot grid work myself, but seem to be getting fairly reliable recovery of sections at present (after quite a steep learning/disappointment curve). I will share my process with you although I am sure there are more experienced users out there that might have more tips and tricks. As far as I'm aware there's no magic trick (I would certainly like to hear it myself if there is), but after a bit of practice, patience and trial and error, it's not too difficult to get sections on the grid without too much destruction.....most of the time (I still have some that get away).

I don't know precisely what you are trying so I will just describe what I do and you can see where it does or doesn't differ from your approach.

1) Cut sections (obviously); I tend to do just enough for 3 or four grids at a time as I find it easier when the surface of the water isn't too crowded (often this is all I need anyway)

2) Stretch/dewrinkle with a heat pen (or chloroform, makes no difference in my hands other than health and safety considerations)

3) Pick up plastic coated slot grid with locking, curved forceps (I use pioloform (0.5%) rather than formvar for my slot grids, although I use formvar for coating of mesh grids for negative staining of viruses. This is historical (ie habit), I don't know whether pioloform is better for slot grids (comments anyone?), but it is what I was shown and it works for me)

4) I hold the forceps holding the grid in my left hand (plastic-coated surface on the top of the grid as I look at it) and submerge the grid under the water away from any sections. Submerging the grid perpendicular to the surface causes least disruption to sections.

5) Holding a "section tickler" (the ubiquitous eyelash glued to the end of a thin wooden dowel) in my right hand, I carefully move and align the section or sections over the surface of the water to the spot above the submerged grid

6) The grid is then slowly pulled out of the water at quite a steep angle (probably } 60ish degrees relative to the surface of the water, but play around to see what's best for you). As the grid emerges from the water the surface tension will tend to push the sections away (as you have noted), but you have to use the eyelash to hold the sections approximately in place, by using either the shaft of the eyelash behind the sections (ie furthest from the grid, pushing) or the tip of the eyelash at the front of the sections (ie nearest to the grid, pulling)). The pioloform surface of the grid is hydrophobic and as it emerges from the water it is quite sticky for the epon section. As the grid is surfacing you need to push or pull (depending whether you are using the eyelash shaft or tip, respectively) an edge or corner of the section to bring it into contact with the pioloform on the surface of the grid. Once in contact it will stick and you then slowly pull out the grid at 60ish degree angle to the surface of the water. The direction of withdrawal from the water should be sympathetic with the alignment of the section rather than the alignment of the slot. By that I mean it is better to have smooth sections that are only partially lined-up with the slot (ie overhanging on either side) rather than sections that are enclosed in the slot but have become creased and wrinkled in the process (from your description, maybe this is where you are going wrong?).

7) Place the grid, still locked in the forceps, carefully onto the bench to dry for a few minutes. The pioloform surface of the grid with the section (hopefully) attached emerges from the water already virtually dry but there is usually a drop of water trapped on the back of the grid. When I have been impatient and tried to dry with fliter paper I have usually poked a hole in the pioloform, so its best to resist! Once this has air dried the grid can be processed further.

Obviously the crucial step is number 6. Another way that I have heard described to me is that you bring the grid out almost perpendicular to the surface of the water and use the eyelash to hook a corner of the section (or ribbon) over the edge of the grid as it surfaces. Once the section is attached to the grid it is carefully withdrawn as above. I have tried this "edge-hooking" way, and (in my limited experience) although it is easier to get the section attached to the grid, I found it more difficult to get the section lined up with the slot. Could be I just need more practice.

Another crucial point in my lab was to have the airflow altered so that there were no draughts affecting the surface of the water. It is difficult enough as it is getting the sections and slot lined up without a mini-maelstrom in your knife boat.

Hope you find something in here that is useful. If there is anything that is still unclear, please don't hesitate to ask.

Good luck

Matthew

Matthew J Hannah PhD
Lead Electron Microscopist
Virus Reference Department
Health Protection Agency
61 Colindale Avenue
London NW9 5EQ
tel: 020 8327 6386

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Email: gburgess-at-dsmanitoba.ca
Name: garry burgess

Organization: Health Sciences Center

Title-Subject: Using Formvar Coated Slotted Grids

Message: I have having HUGE problems getting my ultrathin sections to go
on Formvar Coated grids without a big fight with the sections and the
water in the boat of the diamond knife.

Because of the surface tension of the water, I'm having a big problem
getting the section onto the grid without the making a lot of wrinkles
in the formvar coating, or the section or both.

Is there a trick to this that I don't know about?

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Email: gburgess-at-dsmanitoba.ca
Name: garry burgess

Organization: Health Sciences Center

Title-Subject: Using Formvar Coated Slotted Grids

Message: I have having HUGE problems getting my ultrathin sections to go
on Formvar Coated grids without a big fight with the sections and the
water in the boat of the diamond knife.

Because of the surface tension of the water, I'm having a big problem
getting the section onto the grid without the making a lot of wrinkles
in the formvar coating, or the section or both.

Is there a trick to this that I don't know about?

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{FONT SIZE=3D"4"} {FONT FACE=3D"Lucida Grande"} {SPAN STYLE=3D'font-size:11pt=
'} Gary- {BR}
The trick is to have an absolutely clean water surface and raise the water =
level so it is convex. Obviously you have to stop the sections coming=
when you do this. Then move the sections around with an eyelash- we =
glue ours to a toothpick. You can position the sections anywhere and =
pick them up right on the center of the grid. {BR}
Carol Heckman {BR}
Bowling Green State University {BR}
{BR}
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Hi Garry,

I have not worked with slotted grids, only regular (square) and
hexagonal mesh, but the problem you describe is not uncommon.
If by �fighting� with the sections you mean that they tend to
want to run away from the grid when you are trying to pick them
up, or because you are having trouble getting your grid into the
water, that is because grids tend to be hydrophobic. Two things
that should help:

1) Use a static-free polypropylene specimen cup to condition
your double-distilled water prior to filling the boat. Ted Pella
has them (Static-Free Plastic Cups, 30 ml, Prod. #12901). I'm
pretty sure they ship to Canada. Fill the cup, wait a couple of
minutes, then use a needle syringe to fill your boat with water
from the cup.

2) Glow-discharge (plasma clean) your grids just prior to picking up
sections. This will make your grids hydrophilic. The discharge is
good for about an hour; I usually plasma clean my Formvar+carbon
grids while the ultramicrotome is churning out thin sections.

With conditioned water and plasma cleaned grids, there is very
little resistance at the water's surface when you slide your grid in,
so there will be no wrinkles on the film due to the grid bending. In
my opinion, using static-free water is better than adding Photo-Flo
or solvents like acetone to your boat to reduce surface tension,
which most knife manufacturers advise against anyway.

One other method that was recommended to me is to store Formvar
coated grids in the refrigerator until you are ready to use them;
this supposedly keeps them hydrophilic. I found that did not work
with our Formvar+carbon grids. If you use only plain Formvar film,
storing your grids at 4 degrees C might help.

To avoid wrinkles in the sections, I relax them with chloroform vapor
before picking them up (just a couple of quick waves because you
don't want to over-stretch them either). I use locking tweezers and
release the grids onto filter paper in a Petri dish. I then place
the Petri dish in a 60 degree C oven for 10 to 20 minutes to ensure
good adhesion and fewer sections washed off during post-staining.

Hope this helps.

Cheers,

Gigi K.

Gigi Kemalyan
Nogales Lab Research Technician
Howard Hughes Medical Institute
University of California, Berkeley
742 Stanley Hall
Berkeley, CA 94720-3220
Ph 510-666-3335
Email (work): gigik-at-berkeley.edu
Email (alt.): singinggardenersx2-at-live.com
Website: http://cryoem.berkeley.edu

----------------------------------------------------------------------

Email: gburgess-at-dsmanitoba.ca
Name: garry burgess

Organization: Health Sciences Center

Title-Subject: Using Formvar Coated Slotted Grids

Message: I have having HUGE problems getting my
ultrathin sections to go on Formvar Coated grids without a big
fight with the sections and the water in the boat of the diamond
knife.

Because of the surface tension of the water, I'm having
a big problem getting the section onto the grid without the
making a lot of wrinkles in the formvar coating, or the section
or both.

Is there a trick to this that I don't know about?

----------------------------------------------------------------------

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I see lots of interesting ideas and methods for picking up serial sections onto formvar-coated slotted grids.

As all involve immersing the coated grids in water and picking up the sections from below, I think they will require lots of skill and patience.

A simpler method is to use uncoated slotted grids as pick-up loops. Align the batches of serial sections on the side of the knife boat and cover each row with an uncoated grid similar to the coated ones.

Make sure each grid is completely wet on the bottom surface and then pick it up while keeping it horizontal. The water drop will stay in the slotted hole and the sections will remain on the surface of the drop of water. You will be able to transfer the grid, water drop and sections onto a formvar-coated slotted grid held in a clean pair of forceps. Align the slot in the pick-up grid with the slot on the formvar coated grid and leave to dry slowly. The sections will remain flat and dry down onto the formvar surface.

I think there is a similar version where the grid with the drop of water and sections are placed on formvar sheets created over the holes of a sheet of perspex. Once dry, the grid, with formvar and sections are removed.

Have fun trying this - I promise you it will be easier than picking the sections up from below.

I got first the idea from an old (1970's) paper which I do not remember the reference for (or the authors - maybe it was Galey and Nielssen).

Regards,

Paul.

Paul Webster, Ph.D.
House Research Institute
2100 West Third Street
Los Angeles
CA 90057.

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Hi Paul,

a somewhat similar method was used by Marinozzi to pick up sections and isolate them in a plastic loop. Was quite useful for cytochemical procedures that were aggressive for metals such as for the detection of glyco conjugates using the Thi�ry method. Way back I used the perforations of 35 mm film to transfer sections, that might work for slotted grids as well.

Marinozzi, V.
Cytochimie ultrastructurale du nucleole . RNA et proteines intranucleolaires. J. Ultrastruct. Res. 10, 433 (1964).

Cheers all,

Jan

Jan Leunissen
Aurion

i: www.aurion.nl

On 16/06/2012, at 3:42 PM, pwebster-at-hei.org wrote:

} ----------------------------------------------------------------------------
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} I see lots of interesting ideas and methods for picking up serial sections onto formvar-coated slotted grids.
}
} As all involve immersing the coated grids in water and picking up the sections from below, I think they will require lots of skill and patience.
}
} A simpler method is to use uncoated slotted grids as pick-up loops. Align the batches of serial sections on the side of the knife boat and cover each row with an uncoated grid similar to the coated ones.
}
} Make sure each grid is completely wet on the bottom surface and then pick it up while keeping it horizontal. The water drop will stay in the slotted hole and the sections will remain on the surface of the drop of water. You will be able to transfer the grid, water drop and sections onto a formvar-coated slotted grid held in a clean pair of forceps. Align the slot in the pick-up grid with the slot on the formvar coated grid and leave to dry slowly. The sections will remain flat and dry down onto the formvar surface.
}
} I think there is a similar version where the grid with the drop of water and sections are placed on formvar sheets created over the holes of a sheet of perspex. Once dry, the grid, with formvar and sections are removed.
}
} Have fun trying this - I promise you it will be easier than picking the sections up from below.
}
} I got first the idea from an old (1970's) paper which I do not remember the reference for (or the authors - maybe it was Galey and Nielssen).
}
} Regards,
}
} Paul.
}
} Paul Webster, Ph.D.
} House Research Institute
} 2100 West Third Street
} Los Angeles
} CA 90057.

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Hi Garry,

Reading the other posts reminded me of one more method, using
a "Perfect Loop for ultramicrotomy". EMS sells them and also
has the how-to illustrations on their product page. Not cheap
but it works well, at least for regular grids. Might be worth
a try with your slotted grids.

http://www.emsdiasum.com/microscopy/products/preparation/ultramicrotomy.aspx#70944

Cheers,

Gigi K.

Gigi Kemalyan
Nogales Lab Research Technician
Howard Hughes Medical Institute
University of California, Berkeley
742 Stanley Hall
Berkeley, CA 94720-3220
Ph 510-666-3335
Email (work): gigik-at-berkeley.edu
Email (alt.): singinggardenersx2-at-live.com
Website: http://cryoem.berkeley.edu

---------------------------------------------------------------

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Email: warmundm-at-missouri.edu
Name: Michele Warmund

Organization: University of Missouri

Title-Subject: unidentified cellular material

Message: I have images of epidermal cells with magenta-colored erinea (hairs) of the velvet petiole
gall on black walnut (Juglans nigra) trees. These galls are induced by an eriophyid mite, Aceris
caulis. I suspect that the darkly stained cellular material in the first two images posted at
http://extension.missouri.edu/warmund/ is a phenolic material or tannin. We fixed the tissue in 2%
gluteraldehyde and 2% paraformaldehyde with and without 1% caffiene. Caffiene was suggested to us to
prevent leaching of phenols into the vacuole. The third image is simply a cross section of the
petiole gall with the brightly colored erinea. Could anyone please offer comments, verify, or
suggest what the darkly stained cellular material is? Thank you so much for your consideration of
this matter.
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Email: nelofolha-at-gmail.com
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Title-Subject: microscopy using eyeglasses

Message: Hello,

I am from Portugal and i am thinking to buy a microscope. I am thinking to buy it from internet and
i would ask you one thing is very important for me: I use glasses and i would like to know what
feature i must look to know if i can see throught them with glasses. Could you help me? My mail is:
nelofolha-at-gmail.com

Thank you very much (sorry my English)

Domingos Folha

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Name: Domingos Folha

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Title-Subject: microscopy using eyeglasses

Message: Hello,

I am from Portugal and i am thinking to buy a microscope. I am thinking to buy it from internet and
i would ask you one thing is very important for me: I use glasses and i would like to know what
feature i must look to know if i can see throught them with glasses. Could you help me? My mail is:
nelofolha-at-gmail.com

Thank you very much (sorry my English)

Domingos Folha

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Colleagues

There has been a problem with the Microscopy Listserver automatic archiving system.

I have restored the missing records April 22 -} June 16 .

Archives through the this morning can be retrieved at

http://www.microscopy.com/MicroscopyArchives.html

but automatic daily updates will likely not be fixed until I return at the end of June.

Sorry,

Nestor
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Dear Domingos,

The main thing you need to do is make sure the eyepieces on your
microscope are adjustable, and have a high eyepoint, so you can focus
through them while wearing your glasses.

I also wear glasses, and my eyes are so different that I cannot get both
eyes in focus using only the adjustment in the eyepieces - I have to wear
glasses while using the microscope. As long as the eyepieces are high
eyepoint and adjustable, I have no trouble.

best regards,
Rosemary White

Dr Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
E rosemary.white-at-csiro.au

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Hi Michele,

Those osmiophilic inclusions do look rather characteristic of phenolics -
anthocyanins often stain this way, too. If you took a thin hand-section of
the gall, put on a slide, and observed under a compound microscope, you
could probably get a macro-view of the anthocyanins in these cells.

Couple of examples:
www.plantphysiol.org/content/130/2/561
www.plantphysiol.org/content/158/2/666

cheers,
Rosemary

Dr Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
E rosemary.white-at-csiro.au

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Hello again,

Many people wrote to me off-line to point out that EMS has a "Perfect Loop" that will do the same thing I described for picking up sections.

I agree that the Perfect Loop is useful for picking up sections to place onto a regular grid where orientation is not essential.

However, the original question was for advice on how to pick up sections onto a slotted grid. This is a single slot grid which is longer than it is wide.

Orienting a ribbon of sections onto the formvar-coated slot will be very difficult is using the Perfect Loop, which has no orientation. Using a clean slot grid makes it possible to orientate the sections before they dry down. Maybe there is a market niche for an oval-shaped Perfect Loop!

Sincerely,

Paul Webster.

Los Angeles.

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I used to use the slot grid as the loop, with regular success.
Briefly:
A domino rack was coated with 2.5% formvar in dry chloroform, and the grids were "coated" (to make them sticky) by dipping in 0.5% Formvar and air drying immediately on filter paper.

Put the coated domino rack onto a 60 degree C hotplate.
Line your sections up in the boat, and using the slot grid, dull side down, as your loop, pick up sections from above the water. The sections will ALWAYS be in the hole with this method. Hold the loaded slot grid above a hotplate for a few seconds to flatten the sections, then GENTLY place onto formvar film over a hole in the domino rack. Record position of grid on rack. Allow to dry for at least an hour, allow to cool, store in dessicator overnight, then gently "punch" the grid out of the rack. The hex key tool in the Leica cryo kit is perfect for this.
Stain the Shiny side!

Best regards,
Rosey

Rosey van Driel
Manager, Transmission Electron Microscopy
Institute for Frontier Materials
Geelong Technology Precinct
Deakin University,�
75 Pigdons Rd
Waurn Ponds, Vic 3217,� Australia

Telephone: +61 3 5227 3117 Mobile: 0417399630
Facsimile: 03 5227 1103
Email:rosey.vandriel-at-deakin.edu.au
Website:http://www.deakin.edu.au
Deakin University CRICOS Provider Code 00113B

-----Original Message-----
X-from: PWebster-at-hei.org [mailto:PWebster-at-hei.org]
Sent: Monday, 18 June 2012 3:05 PM
To: Rosey Van Driel

Hello again,

Many people wrote to me off-line to point out that EMS has a "Perfect Loop" that will do the same thing I described for picking up sections.

I agree that the Perfect Loop is useful for picking up sections to place onto a regular grid where orientation is not essential.

However, the original question was for advice on how to pick up sections onto a slotted grid. This is a single slot grid which is longer than it is wide.

Orienting a ribbon of sections onto the formvar-coated slot will be very difficult is using the Perfect Loop, which has no orientation. Using a clean slot grid makes it possible to orientate the sections before they dry down. Maybe there is a market niche for an oval-shaped Perfect Loop!

Sincerely,

Paul Webster.

Los Angeles.

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Hi Garry and all,
Our lab doesn't have a glow discharge unit so we mainly use the
formvar-coated bridge method as described by J. Carter Rowley and
David T. Moran (1975). A Simple Procedure for Mounting Wrinkle-Free
Sections on Formvar-Coated Slot Grids. Ultramicroscopy 1, 151-155.

Where we differ from their method - we submerge the slot grids at an
angle keeping the slot filled with water and then scoot the sections
into the slot with an eyelash tool rather than lower the slot down
onto the sections. The sections will float in a meniscus of water and
after you place the grid down onto the formvar-coated bridge the
sections will dry down onto the formvar. We usually let them dry down
for 2 hrs but that may be overkill. And, we don't try to coat the
slots with carbon after picking up the sections. The formvar alone
seems to give enough support.

Anti-capillary forceps are a huge help when using slot grids and I'm a
big fan of Synaptek 2x1 mm slot grids with dots. I pick up the
sections with the dot side up so I know to stain the grids dot side
down.

Roesy van Driel gave good tips in her reply (dipping the grids prior
to use to make them "sticky"), etc. tho we haven't tried using a
hotplate - maybe the heat and humidity here in Georgia allows us to
skip that step;-)

my best,
Beth

On Jun 14, 2012, at 4:26 PM, zaluzec-at-Microscopy.Com wrote:

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} Email: gburgess-at-dsmanitoba.ca
} Name: garry burgess
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} Title-Subject: Using Formvar Coated Slotted Grids
}
} Message: I have having HUGE problems getting my ultrathin sections
} to go
} on Formvar Coated grids without a big fight with the sections and the
} water in the boat of the diamond knife.
}
} Because of the surface tension of the water, I'm having a big problem
} getting the section onto the grid without the making a lot of wrinkles
} in the formvar coating, or the section or both.
}
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9, 22 -- From beth-at-plantbio.uga.edu Mon Jun 18 11:00:28 2012
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Dear Paul,
Dear all,
only in case anybody would like to see the paper you/Paul cited / tried to cite (see below)
I would like to inform that it is/was:

Galey FR, Nilsson SE.
A new method for transferring sections from the liquid surface of the trough through staining solutions to the supporting film of a grid.
J Ultrastruct Res. 1966 Feb;14(3):405-10.
(for an abstract see also: http://www.sciencedirect.com/science/article/pii/S0022532066800576)

other sources and hints found (Disclaimer: naming of companies etc. without any commercial interest):
see ('Chien' Grids) : http://www.tedpella.com/grids_html/tomography.htm
see (Oval slot, Chien-grid, bottom of webpage): http://www.emsdiasum.com/microscopy/products/grids/embra.aspx

best regards,

Wolfgang MUSS
SALZBURG
Austria

================================================================
Von: PWebster-at-hei.org [mailto:PWebster-at-hei.org]
Gesendet: Samstag, 16. Juni 2012 05:44
An: Mu� Wolfgang
Betreff: [Microscopy] Re: Using Formvar Coated Slotted Grids // collecting sections on slotted grids

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I see lots of interesting ideas and methods for picking up serial sections onto formvar-coated slotted grids.

As all involve immersing the coated grids in water and picking up the sections from below, I think they will require lots of skill and patience.

A simpler method is to use uncoated slotted grids as pick-up loops. Align the batches of serial sections on the side of the knife boat and cover each row with an uncoated grid similar to the coated ones.

Make sure each grid is completely wet on the bottom surface and then pick it up while keeping it horizontal. The water drop will stay in the slotted hole and the sections will remain on the surface of the drop of water. You will be able to transfer the grid, water drop and sections onto a formvar-coated slotted grid held in a clean pair of forceps. Align the slot in the pick-up grid with the slot on the formvar coated grid and leave to dry slowly. The sections will remain flat and dry down onto the formvar surface.

I think there is a similar version where the grid with the drop of water and sections are placed on formvar sheets created over the holes of a sheet of perspex. Once dry, the grid, with formvar and sections are removed.

Have fun trying this - I promise you it will be easier than picking the sections up from below.

I got first the idea from an old (1970's) paper which I do not remember the reference for (or the authors - maybe it was Galey and Nielssen).

Regards,

Paul.

Paul Webster, Ph.D.
House Research Institute
2100 West Third Street
Los Angeles
CA 90057.

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Email: ian.maclaren-at-glasgow.ac.uk
Name: Ian MacLaren

Organization: University of Glasgow

Title-Subject: Readership in Condensed Matter Physics

Message: I would like to draw your attention to a Readership with our
group at the University of Glasgow. Our research focuses on the physics
of materials, with an emphasis on the nanoscale and particular expertise
in electron microscopy. Activities range from experimental and
theoretical studies of fundamental phenomena to developing solutions for
critical issues such as energy, healthcare and information technology.
This research is underpinned by advanced characterisation, modelling and
a long-standing reputation for the development of transmission electron
microscopy techniques.

The closing date for application is 27 June, 2012.

If you know of anyone who may be interested, please direct them to:
http://www22.i-grasp.com/fe/tpl_glasgow01.asp
and enter job reference 001788

Best wishes

Ian MacLaren

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Email: jianzuo-at-uiuc.edu
Name: Jian-Min Zuo

Title-Subject: Joint postdoc position for Advanced Electron Microscopy

Message: Dear Fellow Microscopists,

I am posting this job description, hopefully you can forward this to
interested candidates.

Thank you for your help!

Sincerely

JM Zuo

Electron Diffraction for advanced materials characterization

We are seeking for a highly-qualified and motivated postdoctoral fellow
with an interest in electron nanodiffraction and quantitative electron
microscopy for advanced materials characterization in general. The
position is founded by the French Nanoscience Foundation at Grenoble for
18 months (and certainly a possibility to extend to 2 years). The
successful candidate will be jointly supervised by Dr. Jean-Luc Rouviere
of CEA-Grenoble, France and Prof. J.M. Zuo of Nanoscience Fondation,
Grenoble, France and University of Illinois, Urbana-Champaign, USA.

The candidate will work in the Electron Microscopy Laboratory of the
Institute of Nanoscience and Cryogenic (inac.cea.fr/en/) and the
Nanocharacterisation Center in the Minatec campus (www.minatec.org/).
The laboratory uses a variety of microscopy techniques for atomic and
nanoscale materials characterization with state of art instruments,
including a TITAN Ultimate (50 pm resolution) transmission electron
microscope equipped with monochromator and both image and probe
correctors. Specific project areas include development and applications
of local electron diffraction techniques using coherent electron beams
of sub-nm to tens of nm, combination of diffraction and image
information for quantitative analysis of composition, strain, interfaces
or defects. Materials of interest include semiconductor interfaces,
nanowires and quantum dots.

Grenoble and Minatec, situated in the French Alps, offer a fantastic
quality of life combined with excellent research facilities.

Position Requirements: PhD in materials science, or physics or related
discipline and related scientific experience. Compensations will be
commensurate with research experience. Position can start as soon as
possible.

To be considered, please send CV, brief description of research
interests and contact information of references to either
Dr Jean-Luc ROUVIERE Prof. Jian-Min ZUO
Tel.: +41 4 38 78 50 86 Tel.: +01 217 244 6504
Fax: +41 4 38 78 58 17 Fax: +01 217 333 2736
Email: jean-luc.rouviere-at-cea.fr

or
Email: jianzuo-at-illinois.edu
http://cbed.matse.illinois.edu

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Hi Nicki,
Domino racks are a copolymer coated metal rack with 5mm diameter holes in them. (Available from EMS # 70620. No commercial interest). There is a reference with the product listing on the EMS website.

Best regards,
Rosey

Rosey van Driel
Manager, Transmission Electron Microscopy
Institute for Frontier Materials
Geelong Technology Precinct
Deakin University,�
75 Pigdons Rd
Waurn Ponds, Vic 3217,� Australia

Telephone: +61 3 5227 3117 Mobile: 0417399630
Facsimile: 03 5227 1103
Email:rosey.vandriel-at-deakin.edu.au
Website:http://www.deakin.edu.au
Deakin University CRICOS Provider Code 00113B

==============================Original Headers==============================
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Dear colleagues

I will appreciate if you can distribute this announcement to your
colleagues. Thank you!

Sergei

The Center for Nanophase Materials Sciences (CNMS) at Oak Ridge National
Laboratory (ORNL) is seeking to fill a postdoctoral position in the
field of advanced force-based Scanning Probe Microscopy (SPM). This
program takes advantage of recently developed R&D100 Award winning Band
Excitation Scanning Probe Microscopies, including new spectroscopic
modes for studies of energy dissipation and energy transfer, nanoscale
thermal and electromechanical imaging of a wide range of materials,
including oxides and polymers. This position provides an opportunity to
join an experienced team to develop methods and applications in
directions such as dynamic SPM and new control modes, multifrequency
SPM, and time-resolved measurements. The program will have a strong
component of innovative technique development and application to a broad
range of materials systems. The CNMS is a collaborative nanoscience user
research facility established by the Office of Science, U.S. Department
of Energy.

Qualifications:

This position requires a Ph.D. in Electrical Engineering, Condensed
Matter Physics or Materials Science. The successful applicant should
have a strong background in Matlab and Labview programming,
Field-programmable gate array programming, and control and feedback
systems. Knowledge of scanning probe microscopy is highly beneficial,
but not required. Prior research work must be supported by a strong
record of publications in peer-reviewed journals and presentations at
scientific conferences. The applicant should be motivated, safety
conscious and possess excellent oral and written communication skills.
The applicant must have the ability to work in a team, interact
effectively with colleagues and external collaborators. Demonstrated
ability to communicate in English to an international scientific
audience is essential. Applicants must have received their highest
degree not more than five years prior to the date of application, and
must complete all degree requirements before starting the appointment.

Technical Questions
For more information about this position, please contact Dr. Stephen
Jesse (sjesse-at-ornl.gov {mailto:sjesse-at-ornl.gov} ) and Sergei V. Kalinin
(sergei2-at-ornl.gov {mailto:sergei2-at-ornl.gov} ) and reference the ad number
listed above in the subject line. Applications will be accepted until
the position is filled.

http://orise.orau.gov/science-education/internships-scholarships-fellowships/description.aspx?JobId=10580

==============================Original Headers==============================
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Listers,

It has been brought to our attention that when the announcement went out for
the position of Electron Microscopy, Deputy Core Director at the NIH there
was an omission of "nih" in the address for the contact person. It is also
incorrect on the MSA Jobs site. The correct information should read:

Materials should be sent to Dr. Mathew Daniels c/o Trina Gregory,
Administrative Officer, NHLBI, by email: gregoryp-at-nhlbi.nih.gov
{mailto:gregoryp-at-nhlbi.nih.gov} .

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E � Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}

Opinions and experiences related are those of Pat Connelly and do not
represent the NIH. This message is not confidential and can be freely shared
and reproduced.

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The faculty and staff of the LeRoy Eyring Center encourage attendees of Microscopy & Microanalysis 2012 to visit the Southwestern Center for Aberration Corrected Electron Microscopy, located on ASU's Tempe campus.

Open House hours are:

Sunday, July 29 from 10am - 5pm

Thursday, Aug. 2 from 3pm - 6pm

Friday, Aug. 3 from 10am - 5pm

�

Please RSVP to Beth Crespo� |� csss-at-asu.edu� |� 480-965-4544

for more details:
http://le-csss.asu.edu/Microscopy%20Open%20House%20for%20M%2526M

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Hi All,

Are there any downsides to using non-crystallizing colloidal silica? It sounds like it is just as good as normal colloidal silica -- so why do they still sell normal colloidal silica? How does it avoid crystallizing anyway?

Cheers,

Zack Gainsforth
Space Sciences Laboratory, UC Berkeley
7 Gauss Way
Berkeley, CA 94720
510-642-9733
zackg-at-ssl.berkeley.edu

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The crystallizing type of colloidal silica gives a chemo-mechanical polish
(CMP) to some materials. The most common use for it is for CMP of silicon
based semiconductors. For most materials, the non-crystallizing type works
very well. The two most well-known crystallizing CS are SytonR and
Glanzox {. From what I understand, the non-crystallizing colloidal silica is
pH balanced to avoid the crystallizing condition.

Syton will crystallize if you look at it too long. Glanzox is less so. I
avoid using both of them and will only use the non-crystallizing type.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

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Hi All,

Are there any downsides to using non-crystallizing colloidal silica? It
sounds like it is just as good as normal colloidal silica -- so why do they
still sell normal colloidal silica? How does it avoid crystallizing anyway?

Cheers,

Zack Gainsforth
Space Sciences Laboratory, UC Berkeley
7 Gauss Way
Berkeley, CA 94720
510-642-9733
zackg-at-ssl.berkeley.edu

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Email: rra-at-stowers.org
Name: Rhonda Trimble

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Title-Subject: TOTO

Message: I am just curious, does anybody out there still do a TOTO
method, and if so, what is the exact procedure?

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CellPie is a celluar images analysis tool dedicated for spatial pattern
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Good Morning,

Dear Rhonda,
interesting question but, unfortunately, you did not tell about the purpose of your study (-ies):
TEM or SEM? Immuno-LM?

I am aware of
i) an OTOTO-Method for SEM.
(OTOTO method (= osmium-thiocarbohydrazide-osmium-thiocarbohydrazide-osmium))

ii) TOTO therefore I interpret (as others do) either as
{thiocarbohydrazide-osmium-thiocarbohydrazide-osmium} OR
{Tannic acid - Osmium - Tannic acid - Osmium} ,
the latter in the Literature also sometimes found as GTO-staining {Glutaraldehyde-Tannic acid-osmium} incubation of tissue samples (for SEM as well as TEM).

and iii): ?you are not talking on:
a commercial dye {TOTO(-3)} to stain DNA (red as observed in a fluorescence microscope)? Do you?

Apologize if these questions/statements you'll find silly.

If you want to do OT(OTO) - SEM I would like to refer you to

{The Use of Osmium-Thiocarbohydrazide for Structural Stabilization and Enhancement of Secondary Electron Images in Scanning Electron Microscopy of Pollen (with apparently extensive OTOTO protocol)}
by William F. Chissoea, Edward L. Vezeyb & John J. Skvarlac in
Grana, Volume 34, Issue 5, 1995 pages 317-324
DOI: 10.1080/00173139509429065
FREE ACCESS ARTICLE: http://www.tandfonline.com/doi/abs/10.1080/00173139509429065

==} http://www.ncbi.nlm.nih.gov/pubmed/7310442 (unfortunately no online version available)
Friedman PL, Ellisman MH.
Enhanced visualization of peripheral nerve and sensory receptors in the scanning electron microscope using cryofracture and osmium-thiocarbohydrazide-osmium impregnation. J Neurocytol. 1981 Feb;10(1):111-31.

The Use of Osmium-Thiocarbohydrazide-Osmium (OTO) and Ferrocyanide-reduced Osmium Methods to Enhance Membrane Contrast and Preservation in Cultured Cells
MARK C. WILLINGHAM and ANGELINA V. RUTHERFORD
Journal of Histochemistry and Cytochemistr Vol. 32, No. 4, pp. 455-460, 1984
(Also with extensive recipe-section and protocols. pdf in my files, if you would like to have this one, please feel free to contact me again)

Perhaps also cf. Jongebloed et al, Scanning Microscopy 1997:
http://freedownload.is/pdf/new-ultrastructural-evidence-for-the-existence-of-glycocalyx-on-18210625.html
(but no recipe in there, but another basic reference Jongebloed WL, Kalicharan D., et al. Application of the OTOTO non-coating technique; A comparison of LM, TEM, and SEM.Microscopy & Analysis, 1992, 28; 31-33)
==} better you link to:
www.ecmjournal.org/journal/smi/pdf/smi98-2-10.pdf (more recent version = 1998)of the former)

(I have that paper in my files as .pdf)

Os-Tannic Acid-(UO2-Ac-Lead-)Staining of Ultrathin sections:

Ultrastructural Pathology, 1990, Vol. 14, No. 6 : Pages 519-527
Rapid Contrasting of Extracellular Elements in Thin Sections
Koert P. Dingemans and Marius A. van den Bergh Weerman
(doi: 10.3109/01913129009076139)

Sorry if I can't be of further help unless you tell us something about your spec-prep. parameters,
Good luck,
Best wishes and regards,

Wolfgang MUSS, PhD
SALZBURG, Austria

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} Organization: Stowers Institute
} Title-Subject: TOTO
} Message:

I am just curious, does anybody out there still do a TOTO method, and if so, what is the exact procedure?
}
} Thanks,
} Rhonda Trimble
} Stowers Institute
} rra-at-stowers.org
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Email: serene_ng-at-dsi.a-star.edu.sg
Name: Serene

Organization: DSI

Title-Subject: Magnetic sample (EDX map)

Message: Dear all,

I'm working with magnetic materials for STEM and EDX mapping. However,
sample drift makes it terribly hard to get any useful info from my EDX maps.

Are there adjustments that you can recommend for better mapping conditions?

On the same note, how can I tell if my column/len has been contaminated
with any magnetic material?

Thank you.

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Hi Serene,

Ferromagnetic samples can escape from some specimen holders, it is best
if some kind of a screw-type mechanism holds these samples securely. If
the column is contaminated with magnetic samples you might find problems
correcting image astigmatism or see distortions in diffraction
patterns. IF you have a complete record of the settings of the
alignments (e.g. beam tilt for objective optical axis) from the
installation and perhaps the strength of the objective stigmator, you
can compare these values with those you currently have using a carbon
film test specimen (e.g. combined test specimen with gold islands and
graphitised carbon). If for example the beam tilt is far away from it's
"normal" value you might suspect contamination.

Sample drift is usually associated with thermal equilibrium (or lack of
it rather), since the sample holder is often contracting for some time
after insertion. If the magnetic force is great enough to overcome the
spring return force, that often holds a tilted sample cup against it's
drive mechanism, you might find the whole sample cup is flipped towards
the vertical and any chance of microscopy is lost. If you can reduce
the amount of magnetic material present (e.g. by preparing a FIB
specimen) or dilute the number of particles you should be able to reduce
magnetic distortions and if your sample is loose it might be moving to
escape the substrate.

Finally try to work at the highest voltage possible, assuming your
samples are stable and securely mounted.

Good luck

}
}
}
} Email: serene_ng-at-dsi.a-star.edu.sg
} Name: Serene
}
} Organization: DSI
}
} Title-Subject: Magnetic sample (EDX map)
}
} Message: Dear all,
}
} I'm working with magnetic materials for STEM and EDX mapping. However,
} sample drift makes it terribly hard to get any useful info from my EDX maps.
}
} Are there adjustments that you can recommend for better mapping conditions?
}
}
} On the same note, how can I tell if my column/len has been contaminated
} with any magnetic material?
}
}
} Thank you.
}

--
Robert Keyse
EM Facility
Whitaker Laboratory
5 East Packer Avenue
Bethlehem
PA 18015
USA

Tel. +1 610 758 4283
Fax +1 610 758 4244

==============================Original Headers==============================
12, 24 -- From rok210-at-lehigh.edu Wed Jun 20 13:27:30 2012
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I use co-silica of both types from Allied High Tech and the
crystallizing type from Buehler and Exetec. I also use Glanzox.
All of them seem the same to me in terms of polish quality and
they all make the same huge mess if you don't clean your samples
and wheels properly after use.

On 6/19/2012 6:57 PM, S.Walck-at-cox.net wrote:
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} The crystallizing type of colloidal silica gives a chemo-mechanical polish
} (CMP) to some materials. The most common use for it is for CMP of silicon
} based semiconductors. For most materials, the non-crystallizing type works
} very well. The two most well-known crystallizing CS are SytonR and
} Glanzox {. From what I understand, the non-crystallizing colloidal silica is
} pH balanced to avoid the crystallizing condition.
}
} Syton will crystallize if you look at it too long. Glanzox is less so. I
} avoid using both of them and will only use the non-crystallizing type.
}
} -Scott
}
} Scott D. Walck, Ph.D.
} 5234 Sandalwood PL
} Oceanside, CA 92056
}
} S.Walck-at-cox.net
} SWalck65-at-gmail.com
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} (760) 685-2815 (Cell)
} (760) 758-4384 (Home)
}
}
} -----Original Message-----
} X-from: zackg-at-berkeley.edu [mailto:zackg-at-berkeley.edu]
} Sent: Tuesday, June 19, 2012 2:22 PM
} To: s.walck-at-cox.net
} Subject: [Microscopy] Non crystallizing colloidal silica?
}
}
}
}
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} Hi All,
}
} Are there any downsides to using non-crystallizing colloidal silica? It
} sounds like it is just as good as normal colloidal silica -- so why do they
} still sell normal colloidal silica? How does it avoid crystallizing anyway?
}
} Cheers,
}
} Zack Gainsforth
} Space Sciences Laboratory, UC Berkeley
} 7 Gauss Way
} Berkeley, CA 94720
} 510-642-9733
} zackg-at-ssl.berkeley.edu
}
}
}
}
}
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} 22, 40 -- Subject: RE: [Microscopy] Non crystallizing colloidal silica?
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Email: acalhoun-at-bidmc.harvard.edu
Name: Andrea Calhoun

Organization: Beth Israel Deaconess Medical Center

Title-Subject: Old CryoUltramicrotome Malfunction

Message: My older CryoUltramicrotome is not sensing the that the LN2
dewar is full. Any ideas as to what might be broken? I'm guessing its a
temperature sensor, but I have no idea where its located. This machine
is not under service contract, so it would be great if there was a way I
could fix it myself!

If its something I can't fix on my own, does anyone know an independent
company that would service old ultramicrotomes in the Boston area?

Any info would be great!

-Andrea

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Email: jill.verlander-at-medicine.ufl.edu
Name: Jill Verlander

Organization: University of Florida

Title-Subject: LKB 7801B knifebreaker servicing

Message: We have a couple of LKB knifebreakers and are having trouble
getting good knives from them. We are considering having one of them
serviced/overhauled.

Does anyone have experience, good or bad, in doing this or can anyone
recommend a company to do it?

Many thanks,

Jill

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When we had this problem in a Leica cryostage (oldish model) on an Ultracut E, it was the tube that goes down into the dewar - this contains the temperature sensors. Inside the tube is a long plastic (?) probe with a series of connected diodes (I think, can't quite remember) which close or open depending on the temperature. If one of these isn't working, the LN2 above that level will not be sensed. (I don't think they're in series like old Xmas lights that died if just one of them went out.) Our electronics workshop made a whole new probe plus diodes because it was clear the whole thing was on its last legs. We did have a spare at one stage, but I think we gave it away when we donated the cryo unit (we replaced it with an RMS cryostage - very good and reliable, no commercial interest, etc).

So, you can make it yourself, jsut need to wire it in correclty.

cheers,
Rosemary

Dr Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
________________________________________
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To: White, Rosemary (PI, Black Mountain)

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Email: acalhoun-at-bidmc.harvard.edu
Name: Andrea Calhoun

Organization: Beth Israel Deaconess Medical Center

Title-Subject: Old CryoUltramicrotome Malfunction

Message: My older CryoUltramicrotome is not sensing the that the LN2
dewar is full. Any ideas as to what might be broken? I'm guessing its a
temperature sensor, but I have no idea where its located. This machine
is not under service contract, so it would be great if there was a way I
could fix it myself!

If its something I can't fix on my own, does anyone know an independent
company that would service old ultramicrotomes in the Boston area?

Any info would be great!

-Andrea

Login Host: 134.174.110.7
Listserver Email Form V - 20120416
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Hello Jill,
I had my LKB 2178 glass knife maker refurbished by Ted Pella and I am very happy with the results. Their phone is 530-243-2200.

Stay safe..........
Frank
PS I don't work for them, but I do buy from them.

---------------------------------------------------------------------------

Email: jill.verlander-at-medicine.ufl.edu
Name: Jill Verlander

Organization: University of Florida

Title-Subject: LKB 7801B knifebreaker servicing

Message: We have a couple of LKB knifebreakers and are having trouble
getting good knives from them. We are considering having one of them
serviced/overhauled.

Does anyone have experience, good or bad, in doing this or can anyone
recommend a company to do it?

Many thanks,

Jill

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(I've already replied, but other voices would be good. Phil)

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} realname - Julia Dart
} Email - dart1ja-at-cmich.edu
} ORGANIZATION - Central Michigan University
} EDUCATION - Undergraduate College
} LOCATION - Michigan
} QUESTION - I'm going to be a senior at my university next fall and
} am currently a major in Biology: Microscopy. I haven't taken an
} official microscope class yet, since I switched my major just
} recently, so I don't know what really to expect of these type of
} courses. What is some good advice or recommendations in order to
} succeed in a microscopy course? Also, how would I go about looking
} for internships or educational opportunities that I could pursue to
} further my microscopy experience? Thanks!

--

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5, 24 -- From oshel1pe-at-cmich.edu Fri Jun 22 09:23:22 2012
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Thanks Bill for the interest :o)

I did try "wet" sieving with ETOH, which certainly worked better than
agitated dry sieving. However, the smallest of particles for both the
48um sieve and 15um sieve still remained not passed, which is the problem
when I want to quantify the minerals in each size and with the
quantification being via area in 2d section (SEM).

The study is also about any fractionation for different minerals with
respect to size -- e.g., as if lower density minerals might be smaller
than more dense minerals, so I expect size fractions to differ for
different minerals.

I guess I'm a bit surprised I received no responses from users who may
have experience with "jet" sieves. These use agitated air flow together
with suction to deaggregate and agitate particles while trying to suck
them through the sieve. I don't know how well these might work for
particles this size �K after all the smallest of sieves (e.g., 20um) do not
have a lot of open area.

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Genuinely, Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
cogito ergo ZzoooomM

On 12-06-14 10:25 PM, "Bill & Sue Tivol" {wtivol-at-sbcglobal.net} wrote:

} Dear Michael,
} It occurred to me that sieving might go faster if you suspend your
} particles in a less viscous solvent, such as ethanol (other organics
} or freons may also work, but they would definitely require you to work
} in a hood, and they might evaporate too quickly). Agitating the
} sieving apparatus while separating could prevent small particles from
} hanging up on the cloth could improve your yield. BTW, do each of the
} minerals yield the same fraction during the process, or does one have
} a higher yield than another?
} Yours,
} Bill
}
} On Jun 14, 2012, at 4:32 AM, mshaffer-at-mun.ca wrote:
}
} }
} } Thanks Stephane ?
} }
} } Others had suggested sediment rates as well. However, there exists
} } relatively large density differences between the minerals in these
} } powdered samples, the most important of which are quartz and
} } hematite/magnetite, and carbonates between. Subsequent analysis of
} } the
} } modes would depend on there being absolutely no partitioning of
} } minerals
} } relative to size.
} }
} } Still investigating :o)
} }
} } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
} } Genuinely, Michael Shaffer
} } SEM-MLA Research Coordinator
} } Bruneau Centre for Research & Innovation
} } Memorial University
} } cogito ergo ZzoooomM
} }
} }
} }
} } On 12-06-13 5:28 AM, "Stephane Nizet" {nizets2-at-yahoo.com} wrote:
} }
} } } Hi Michael!
} } }
} } } I don't know jet sieving but you are clearly in a delicate zone
} } } (between
} } } 100 and 10�gm) :-)
} } } Particules bigger than 10�gm usually sediment fast in water (of
} } } course it
} } } depends on the density), so to select particles under this size you
} } } just
} } } suspend the powder in water and leave the suspension for 5 min on the
} } } bench and then pour the supernatant. Repeat the process 3x and the
} } } results should be quite good.
} } } For even smaller particles (sub-micro) centrifugation works well.
} } } I suppose it may be possible to use higher density liquids (than
} } } water)
} } } in order to select bigger particles but I never experienced that.
} } } Best regards,
} } }
} } } Stephane
} } }
} } }
} } }
} } } ----- Original Message -----
} } } From: "mshaffer-at-mun.ca" {mshaffer-at-mun.ca}
} } } To: nizets2-at-yahoo.com
} } } Cc:
} } } Sent: Tuesday, June 12, 2012 6:15 PM
} } } Subject: [Microscopy] separating the smallest of particles
} } }
} } }
} } }
} } }
} } }
} } } ------------------------------------------------------------------------
} } } --
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} } } The Microscopy ListServer -- CoSponsor: The Microscopy Society of
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} } } To Subscribe/Unsubscribe --
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} } } --
} } } --
} } }
} } } I have a size fraction of {75um rock particles that I need to further
} } } separate into } 45um, } 15um & {15um. I've tried wet sieving with
} } } precision
} } } fabric, but because the 15um sieve has only 10% open space it took
} } } much
} } } longer than anticipated, and still did not pass more than 40-50% of
} } } that
} } } size. I'm looking into jet sieving, but thought I'd ask this group
} } } about
} } } their experience with wet sieving, jet sieving, and other possible
} } } methods(?)
} } }
} } } TIA :o)
}
}
}
}

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Title-Subject: Microwave oven in biological sample preparation for the TEM

Message: Any one have experience in use of Microwave oven in biological
sample preparation for the TEM.
Any comment about microwave based sample preparation. (We are dealing
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Hello Ravindra,
We have used a microwave for processing clinical samples for many years. I
would not process any other way. It has cut our turn around time from a week
down to 1-2 days! Native Kidney biopsies are completed (including scoping)
in 1 day. We have had no problems or issues with the processing.
The major advantage I see is the fact that infiltration of the resin takes a
total of 12 min! 3 min in 1:1, 3 min in 3:1 and 3 min twice in 100% resin.
We do not polymerize epoxy in the microwave--just not practical for our
purposes. However, we process LR White sample for immuno (we also do
research for campus PI's) in the microwave and I find that polymerization of
the LR white in the microwave makes trimming and cutting sections from the
block so much better.
Hope this helps.
Pat Kysar, C.E.M.T.
University of California, Davis
School of Medicine
Dept of Pathology and Laboratory Medicine

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Hi Folks

I am looking for a contract services lab that can perform Active Voltage Contrast.

We have some scribe-line via chain structures, fabricated in tungsten, with 2 micron by 2 micron terminal pads.
The over-all size of the test structure is 40 microns by 80 microns.

This far, this structure has resisted our advances using passive voltage contrast.

any leads

thanks so much

bryan from Sunnyvale

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Name: Ravindra Thakkar

Organization: NIMHANS

Title-Subject: Microwave oven in biological sample preparation for the TEM

Message: Any one have experience in use of Microwave oven in biological sample preparation for the TEM.
Any comment about microwave based sample preparation. (We are dealing with ultrastructural medical diagnostic using TEM).

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Brian,

There are two tricks that I have for stubborn passive voltage contrast
samples. One is for overall floating structures where whole chain gets
dark in a FIB - use black "Micron" 005 pen from Sakura brand to ground
one end of the chain - these pens have fairly conductive trace with
width about 200um and are great for drawing long grounding lines under
microscope - final connection to chain is by FIB metal. Another trick is
for chains with "soft" disconnect - use "HUGE" beam currents, up to nA
range, to get enough voltage drop to see location of the defect.

If you have a FIB with vented chamber (a-la FIB200 and all the later
variants) then with small hardware hack and a bit of sample prep effort
you can start doing active voltage contrast in about an hour.

Disconnect stage current cable from the sample holder (it is fairly
useless for the originally-intended purpose anyway) and hard-ground
sample holder to the stage. Use the disconnected cable and its
feed-through to bring in voltage from regulated DC power supply (+/- 10V
range) - your FIB is ready. For the sample preparation - attach short
piece of a bonding wire to the piece of wire-wrap wire, attach wire-wrap
wire to the sample by a drop of epoxy, bend bonding wire close to your
chain in question, trim end of the bonding wire with ophthalmic scalpel,
and fix it with drop of Ted Pelllas colloidal silver - all this takes
about 30min. under inspection microscope. Almost done: load sample into
your FIB, attache wire-wrap wire to the and of the freed stage-current
cable, pump the chamber, make final connection of the chain to the
silver dot at the end of the bonding wire, and apply voltage from the
power supply :)

If for any reason none of the above works - any lab that has in-situ
manipulator on the FIB should be able to do active VC for... EAG in
Sannyvale, or maybe even one of Universities next door?

Cheers :)
Valery

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 6/22/2012 6:49 PM, Bryan.Tracy-at-spansion.com wrote:
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} Hi Folks
}
} I am looking for a contract services lab that can perform Active Voltage Contrast.
}
} We have some scribe-line via chain structures, fabricated in tungsten, with 2 micron by 2 micron terminal pads.
} The over-all size of the test structure is 40 microns by 80 microns.
}
} This far, this structure has resisted our advances using passive voltage contrast.
}
} any leads
}
}
} thanks so much
}
} bryan from Sunnyvale
}
}
}
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I can almost copy the e-mail from Pat Kysar (UC Davis), with our experience on microwave processing, with some minor modifications:
we process kidney samples regularly, since } 2 years now, and I very much prefer these samples against conventional RT processing (i.e. processing for many long hours with too many and long steps of dehydration and infiltration/ embedding).
turn around times: from 4 days (incl polymerization), down to 4 hours (again incl. infiltration and polymerization of Epon! works great!), resulting in specimen which can be sectioned instantly. Here, we continue on the next day, with semithin+LM, trimming and sectioning for TEM. Yes, you can in fact do this on the very same day, if you are focussed on a very special aspect, only. If you aim to do a more detailed visualization of details of kidney ultrastructure, I would argue that you may need one to even a few days on the TEM, alone. (note: here, we have time slots of 3 hours only, on the TEM, which need to be booked in advance, due to a tight scheduling scheme of TEM time).
Nice to read that LR White can be processed in the microwave, too - which conditions are used for infiltration+ polymerization of LR White?
BTW: we used kidney samples - processed in the microwave with little OsO4, 0.1% - and Epon embedding also for immuno-labelling - and it is worth trying: it works! (depending on the initial processing / perfusion fixation of the kidney, and the amount and kind of antigen present in kidney).
We have to restrict our experience to microwave processing in the AMW (and we think this is important to note) - and another note added: we are just a satisfied customer (Leica), no financial interest.
kind regards,
Reinhard

--
Prof. Dr. Reinhard Rachel
University of Regensburg
Centre for EM / Anatomy
Faculty of Biology & Preclin. Med.
Universitaetsstrasse 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-regensburg.de
office: VKL 3.1.29

next microscopy conferences:
http://www.emc2012.org.uk/
EMC 2012 in Manchester, UK
http://www.mc2013.de/
MC2013 in Regensburg, Germany

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Hi Ravi,

Please start here: http://www.tedpella.com/34700_html/ref.htm

Google: "microwave processing transmission electron microscopy"

Cheers,

Fred Monson

http://cmirt.wcupa.edu

________________________________________
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Email: ravi.thakkar369-at-gmail.com
Name: Ravindra Thakkar

Organization: NIMHANS

Title-Subject: Microwave oven in biological sample preparation for the TEM

Message: Any one have experience in use of Microwave oven in biological
sample preparation for the TEM.
Any comment about microwave based sample preparation. (We are dealing
with ultrastructural medical diagnostic using TEM).

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18, 40 -- From FMonson-at-wcupa.edu Mon Jun 25 06:54:41 2012
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18, 40 -- From: "Monson, Frederick" {FMonson-at-wcupa.edu}
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18, 40 -- CC: "ravi.thakkar369-at-gmail.com" {ravi.thakkar369-at-gmail.com}
18, 40 -- Subject: RE: [Microscopy] via-WWW:Microwave oven in biological sample
18, 40 -- preparation for the TEM
18, 40 -- Thread-Topic: [Microscopy] via-WWW:Microwave oven in biological sample
18, 40 -- preparation for the TEM
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18, 40 -- preparation for the TEM
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Microscopy Lab Technician

This position is within the Discovery Research department at Dow AgroSciences in Indianapolis, IN. The primary purpose of this job is to provide support for a wide array of microscopy-related projects that come through the Microscopy and Imaging Lab at DAS Indy.

Some example tasks that would be performed in this role would be:

* Operation of upright, inverted, and stereomicroscopes and training other users on the use of these microscopes
* Preparation of samples for sectioning, including dissection, fixation, dehydration, and embedding of samples in wax and plastic
* Sectioning, mounting, staining, observing, and photographing samples
* Performing immunolocalizations, in situ hybridizations, and other cytochemical labeling procedures
* Imaging samples by SEM, including specimen preparation by critical point drying and sputter coating
* Operation of a laser scanning confocal microscope, both alone and as technical support in a group setting
* Operation of an automated immunostainer
* Maintaining the microscope rooms and wet lab areas
* Ordering and stocking necessary supplies and reagents

If interested, see the full job description and apply at the following location:

https://dow.taleo.net/careersection/10060/jobdetail.ftl?job=1205054

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Email: jinsong-wu-at-northwestern.edu
Name: Jinsong Wu

Organization: Northwestern University

Title-Subject: Senior Research Associates and Postdoctoral Fellows at Northwestern University

Message: Senior Research Associates and Postdoctoral Fellows

Analytical S/TEM of Nanostructured Materials
In-Situ S/TEM of Dynamic Phenomena in Materials
Cryo-Bio Electron Microscopy of Soft/Hybrid Structures
Department Materials Science & Engineering
International Institute of Nanotechnology (IIN)
The NUANCE Center
Northwestern University, Evanston, IL 60208

Several research positions at the level of postdoctoral scholar or senior research associate are
immediately available at Northwestern University in the broad areas of analytical, in-situ and
cryo-bio S/TEM of materials and soft/hybrid nanostructures.
Two positions are focused on projects related to nanostructured materials, particularly
thermoelectrics and related semiconducting systems. Both require extensive expertise in HRTEM
STEM/HAADF, defect analysis, diffraction, EELS/EDS and tomography, while one is inclined more
towards in-situ microscopy covering thermal, fluidic-cell and related measurements.
Another position is geared towards cryo-bio S/TEM imaging/analysis of assembly/organization of soft
and hybrid nanostructures such as patterned DNA/lipids, nanoparticle-DNA/protein assemblies, etc.
These positions are a part of extensive cross-disciplinary and collaborative materials research
activities at Northwestern. Thus, the candidates have ample opportunities to learn and contribute to
analytical/in-situ and cryo-EM in materials science, bio- and nano-technologies.
Northwesterns NUANCE center (http://www.nuance.northwestern.edu/epic) and sister facilities are
well equipped with modern S/TEMs, SEMs and extensive specimen preparation capabilities; ranging from
focused ion beam (FIB) to cryo-transfer with LN2 stages.
The positions require PhD in physical/biological sciences/engineering. Respective experience in
analytical, in-situ and/or cryo-EM techniques and computation/simulations is essential. The
positions are available immediately for at least two years, with possibility for mutual extension.
Salary and compensation would commensurate with experience, up to: $ 55,000-$65,000 per year. Select
candidates are eligible for the IIN Postdoctoral Fellowships, which offer additional significant
compensation on top of the core stipend.

Please indicate position preference; and forward your CV and three references to:
Professor Vinayak P. Dravid
Materials Science & Engineering
Director, NUANCE Center
Northwestern University,
Evanston, IL 60208, USA
Ph.: (847) 467-1363, Fax: (847) 467-3573
E-mail: v-dravid-at-northwestern.edu
URLs: http://vpd.ms.northwestern.edu http://www.nuance.northwestern.edu

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Email: germpore-at-sonic.net
Name: Peter G. Werner

Organization: Ohlone College

Title-Subject: Vacuum pump inlet filter periodic cleaning?

Message: I've recently become the proud new administrator of a Hitachi SU-1500 SEM system. I come
from a background mainly in optical microscopy, and while imaging on an SEM has been straightforward
enough (OK, not really, but not too far out of my experience), it is the first time I've maintained
an instrument with any kind of vacuum system.

I was told by the Hitachi techs to periodically change the oil in the rotary vane vacuum pumps that
power the evacuation system. That's also pretty straightforward (hardly different from changing oil
in a car, really). Consulting the vacuum pump manual for the procedure on this, though, I also
notice that it also recommends cleaning the vacuum inlet filter about twice a year. That raises some
questions.

Is this a point of maintenance that actually needs to be carried out when the pump is fully attached
to a vacuum system, as opposed to one in which the intake valve is continually taking in air from
the surrounding environment (including any dust) rather than drawing out air from inside the EM
system? And if so, what are the conditions under which it would be OK to disconnect the vacuum line
from the EM to the pump? The default state is for the EM chamber and column to be evacuated,
including when the system, including the pumps, are turned off. Would disconnecting the line
backfill the EM with air with potentially bad consequences? Alternately, if I have the EM in Air
(nonevacuated) mode, then the pumps are by default running. I can unplug the power cords one pump at
a time to work on them that would seem to be OK, but they normally both run even in Air mode, so
Im not sure if I should be taking them offline in that mode either. Dilemmas!

Unfortunately, the SEM manuals coverage of the vacuum system is not particularly good, so it leaves
a lot of open questions. Ill run these questions by the Hitachi tech next time I talk with her, but
was wondering how other users treated this particular point of maintenance, or whether you bothered
with it at all.

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Can anyone tell me about the facilities for SEM at Selcuk University? There is a post-doc here who just got introduced to SEM at our materials lab. He will be returning to Turkey soon and it seems he would like to continue to use SEM for characterization. I'd like to find him a contact, if possible.

Warren Straszheim

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Hi

I have to say I am very puzzled because I have never heard of an inlet
filter on an electron microscope??!! Are you sure they are not talking
about the exhaust filter, which I agree should be maintained or replaced
every 6 months? If the exhaust simply ends in a mushroom shaped unit I
would very strongly suggest that you replace it with a good quality filter
as offered in EM accessory manufacturers catalogues.

The pump should be disconnected electrically, switched off, to change the
fluid, but you do not have to switch it off to change the exhaust filter.
But I suggest you do switch off to change filter so that you will not be
exposed to the nasty exhaust fumes.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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Email: germpore-at-sonic.net
Name: Peter G. Werner

Organization: Ohlone College

Title-Subject: Vacuum pump inlet filter periodic cleaning?

Message: I've recently become the proud new administrator of a Hitachi
SU-1500 SEM system. I come from a background mainly in optical microscopy,
and while imaging on an SEM has been straightforward enough (OK, not really,
but not too far out of my experience), it is the first time I've maintained
an instrument with any kind of vacuum system.

I was told by the Hitachi techs to periodically change the oil in the rotary
vane vacuum pumps that power the evacuation system. That's also pretty
straightforward (hardly different from changing oil in a car, really).
Consulting the vacuum pump manual for the procedure on this, though, I also
notice that it also recommends cleaning the vacuum inlet filter about twice
a year. That raises some questions.

Is this a point of maintenance that actually needs to be carried out when
the pump is fully attached to a vacuum system, as opposed to one in which
the intake valve is continually taking in air from the surrounding
environment (including any dust) rather than drawing out air from inside the
EM system? And if so, what are the conditions under which it would be OK to
disconnect the vacuum line from the EM to the pump? The default state is for
the EM chamber and column to be evacuated, including when the system,
including the pumps, are turned off. Would disconnecting the line backfill
the EM with air with potentially bad consequences? Alternately, if I have
the EM in Air
(nonevacuated) mode, then the pumps are by default running. I can unplug the
power cords one pump at a time to work on them that would seem to be OK,
but they normally both run even in Air mode, so Im not sure if I should be
taking them offline in that mode either. Dilemmas!

Unfortunately, the SEM manuals coverage of the vacuum system is not
particularly good, so it leaves a lot of open questions. Ill run these
questions by the Hitachi tech next time I talk with her, but was wondering
how other users treated this particular point of maintenance, or whether you
bothered with it at all.

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I've never seen an inlet filter on a pump, rather I've encountered
such filters on the microscope air inlets. Even older TEMs (Hitachi
HU-11 and H500) had an air inlet desiccator filter. Basically, it was
Drierite or silica gel granules with a large piece of cotton wadding
to prevent loose powder from entering the microscope. We never had one
on our SEMs (Hitachi S570 and 2460), though we did hook up the air
inlet on the 570 to a tank of dry nitrogen gas.

Our most recent SEM, an FEI 450FE, came with a HEPA-like, micro-filter
on the air inlet to prevent the intake of particulates into the EM.
They are basically replaced once or twice a year and the old ones
discarded. No filter on the pump inlet, though.

John J. Bozzola, Ph.D., Professor
Truly, Gratefully Retired Director of IMAGE
Integrated Microscopy & Graphics Expertise
Southern Illinois University
Carbondale, IL 62901

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Actually, I have seen them used extensively on Leybold roughing pumps,
only on the exhaust side. It's a stainless steel trap integrated into
an O ring assembly for a KF25 flange. The trap sits inside the pump and
filters out any large chunks of debris that may fall out of the exhaust
filter. Anyone who wants to see a pic, just email me.

Gary Easton
Scanners Corp.
SEM Service
410-857-7633

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*
***

**Announcing the 2012 CNMS User Meeting*
{http://www.cnms.ornl.gov/workshops/2012/announcement.shtm} *

*Friday, September 14, 2012*
at
Oak Ridge National Laboratory
Oak Ridge, Tennessee

*Click **/here/* {https://register.ornl.gov/2012/cnms/} *//**to
**REGISTER**: */*Deadline: August 31, 2012*/**

*Featuring Workshops*:

�Scanning Probe Microscopy for Energy Applications
{http://www.cnms.ornl.gov/workshops/2012/SPM_EnergyWorkshop2012.pdf} ,
/September 11-13, 2012 /(begins 1pm on the Sept. 11)(abstract submission
{https://register.ornl.gov/2012/cnms/poster_abstract.cfm} )

�Transmission Electron Microscopy for Soft Materials
{http://www.cnms.ornl.gov/workshops/2012/TransElectronMicroscope.shtm} ,
/September 12-13, 2012/

�Second Photovoltaics School
{http://www.cnms.ornl.gov/workshops/2012/Photovoltaics_School.shtm} (Photovoltaics
from Fundamentals to Applications),/September 13, 2012/

*CALL FOR ABSTRACTS*
{http://www.cnms.ornl.gov/workshops/2012/AbstractCall.shtm} /
//deadline August 1, 2012/- for attendees wishing to present their
research at the User Meeting

------------------------------------------------------------------------

The Center for Nanophase Materials Sciences (CNMS)
{http://www.cnms.ornl.gov/} at Oak Ridge National Laboratory is pleased
to announce plans for the 2012 CNMS User Meeting, *to be held on
September 14* at Oak Ridge National Laboratory in Oak Ridge, Tennessee.
Featured workshops will begin earlier in the week.

The annual user meeting combines oral presentations, poster sessions,
workshops and tutorials into a compact program designed to illuminate
the frontiers of nanoscience research and acquaint researchers with the
scientific resources for nanoscience that this user facility offers.

The 2012 User Meeting will feature:

* *Invited plenary speakers*presenting their vision of nanoscience
research challenges and opportunities:
- John Anthony {https://chem.as.uky.edu/users/anthony} , /University
of Kentucky/
- Walter DeHeer
{https://www.physics.gatech.edu/user/walter-de-heer} , /Georgia Tech///

* *Invited user presentations *by users of CNMS facilities:
- William Butler {http://mint.ua.edu/faculty_member/butler-bill/} ,
/University of Alabama
/- J. Todd Hasting {http://www.engr.uky.edu/%7Ehastings/} s,
/University of Kentucky/
/- /Jon-Paul Maria {http://www.mse.ncsu.edu/profile/jpmaria} , /North
Carolina State University/
- Andrew Rinzler {http://www.phys.ufl.edu/faculty/rinzler.shtml} ,
/University of Florida/
/- /Donghui Zhang
{http://chemistry.lsu.edu/site/People/Faculty/Donghui%20Zhang/item1111.html} ,
/Louisiana State University/
-Qiming Zhang
{http://www.ee.psu.edu/Directory/FacultyInfo/Zhang/ZhangProfilePage.aspx} ,
/Pennsylvania State University

/
* *Contributed user talks *to be selected from submitted abstracts
{https://register.ornl.gov/2012/cnms/poster_abstract.cfm} .

* The */RidgeDance NanoScience Film Festival/*
{http://cnms.ornl.gov/workshops/2012/film_festival.shtm} will be back
by popular demand on Thursday evening as well. Attendees are invited
to submit video clips or still images of their work. Contact Chris
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* Complete information on *how to become a user*.
* *User Proposal Planning*- Ample opportunity to meet with facility
staff and other researchers to discuss and plan for submission of
user proposals.
* *User Group business meeting*will provide opportunities for the user
community at-large to provide input on planning and future facility
developments.
* *User Research Posters*: Current facility users and prospective
users will present results of their research and share their
experiences at the user facilities. The User Poster Session on
Thursday evening will provide a great opportunity for grad students
and postdocs to present and discuss their research with colleagues.
Register your poster /here
{https://register.ornl.gov/2012/cnms/poster_abstract.cfm} /, or view
Call for Abstracts
{http://www.cnms.ornl.gov/workshops/2012/AbstractCall.shtm} .

*Advance Registration**is required to attend*this user meeting because
of access control at Oak Ridge National Laboratory, a U.S. Department of
Energy facility (*Deadline*: *August 31, 2012*).On-site registration
will not be available. All attendees are required to register and
provide their name, affiliation, and contact information. Those without
an ORNL badge will also be asked for date and place of birth and
citizenship.

Access to ORNL is for the purpose of attending this user meeting only
and does not enable access to other ORNL facilities or laboratories.
Access to ORNL requires photo identification; foreign nationals will
need to bring their valid passport and visa.

The CNMS user facility is supported by the U.S. Department of Energy�s
Office of Basic Energy Sciences {http://science.energy.gov/bes/} to
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We hope to see you here in September!

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I believe Peter is referring to the various forms of molecular sieves (might
be zeolites or steel or copper wool) that are usually placed near the inlet
of the roughing pump to limit backstreaming of pump oil.

Some of the units are sealed and considered throw-aways, although one can
flush them with acetone and then thoroughly dry them, preferably under
vacuum and heat, before putting them back in the vacuum line. Others
(notably Edwards and M. E. Taylor) can be opened and the contents replaced.

I don't know if his system has a turbo pump or an oil diffusion pump, but
either way, my suggestion would be to shut down completely before opening
any roughing lines.

With isolation valves, the high vac pumps can be left for short periods
without backing (if the vacuum system is tight) but since Peter is new to
vacuum systems, Murphy is watching, and "everything takes longer than it
takes", I would strongly suggest a complete shut down.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
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Bridgton, ME 04009
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kenconverse-at-qualityimages.biz
qualityimages.biz

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Email: germpore-at-sonic.net
Name: Peter G. Werner

Organization: Ohlone College

Title-Subject: Vacuum pump inlet filter periodic cleaning?

Message: I've recently become the proud new administrator of a Hitachi
SU-1500 SEM system. I come
from a background mainly in optical microscopy, and while imaging on an SEM
has been straightforward
enough (OK, not really, but not too far out of my experience), it is the
first time I've maintained
an instrument with any kind of vacuum system.

I was told by the Hitachi techs to periodically change the oil in the rotary
vane vacuum pumps that
power the evacuation system. That's also pretty straightforward (hardly
different from changing oil
in a car, really). Consulting the vacuum pump manual for the procedure on
this, though, I also
notice that it also recommends cleaning the vacuum inlet filter about twice
a year. That raises some
questions.

Is this a point of maintenance that actually needs to be carried out when
the pump is fully attached
to a vacuum system, as opposed to one in which the intake valve is
continually taking in air from
the surrounding environment (including any dust) rather than drawing out air
from inside the EM
system? And if so, what are the conditions under which it would be OK to
disconnect the vacuum line
from the EM to the pump? The default state is for the EM chamber and column
to be evacuated,
including when the system, including the pumps, are turned off. Would
disconnecting the line
backfill the EM with air with potentially bad consequences? Alternately, if
I have the EM in Air
(nonevacuated) mode, then the pumps are by default running. I can unplug the
power cords one pump at
a time to work on them that would seem to be OK, but they normally both
run even in Air mode, so
Im not sure if I should be taking them offline in that mode either.
Dilemmas!

Unfortunately, the SEM manuals coverage of the vacuum system is not
particularly good, so it leaves
a lot of open questions. Ill run these questions by the Hitachi tech next
time I talk with her, but
was wondering how other users treated this particular point of maintenance,
or whether you bothered
with it at all.

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Email: jwoperei-at-smith.edu
Name: Judith Wopereis.

Organization: Smith College

Title-Subject: zebrafish sample for TEM

Message: Hi,

I am looking for a resin block with a `48h zebrafish embryo (or part of the embryo) embedded in it.
I'll be demonstrating the TEM to a group of high school teachers as part of an outreach program that
is focused on zebrafish development. The demonstration will take place in about two weeks.

You can contact me directly if you have a spare block available for me to use.

Your help will be greatly appreciated.

Best,
Judith Wopereis.

Smith College.
Center for Microscopy and Imaging.

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"Philip Oshel" writes:

} First, do you have a service contract on the SEM?
} If yes, then changing the rotary pump oil and any
} filters is (should be) part of the regular
} preventive maintenance, which is part of the
} contract. (At least, it always has been on the
} contracts I've had, including Hitachi.) If you
} only get one P.M./year, then you may have to do
} one change at 6 months. If this is the case, the
} service engineer should have gone over how to do
} this with you at the installation.
} Don't be shy about emailing your service tech and
} asking. The Hitachi folks I deal with (and have
} dealt with elsewhere) are happy to answer
} questions. You don't have to wait.

Interesting! Because we do have a one PM/year contract, which they're due
to come out and take care of soon, but it was the Hitachi tech who came
out on a different matter (misaligned aperture) who told me I really
needed to change the oil! Probably because just looking at the oil level
in one of the pumps, it's clearly overdue. We hadn't had a tech on our EM
in at least 6 months before I started, so I'm dealing with a bit of
deferred maintenance, plus no notes from my predecessor on what the
maintenance needs are for the system.

} Second, when you shut down the vacuum system, the
} valves close to maintain a static vacuum. This
} allows you to do things like change pump oil,
} etc. There is no need to disconnect vacuum lines.
}
} For details of changing the pump oil and filters,
} you should have a separate manual from the pump
} manufacturer (probably Edwards, these days). If
} you don't, you can go to the company's website
} and download or ask for the manual for your model
} pump.

Did that already; that's where I saw the maintenance note for air filter
cleaning. There's no mention in the Hitachi manual. The pump is a Varian
(now Agilent) DS202. I also downloaded the manual for the Edwards oil mist
filter to see about what filters needed to be replaced there and how
often. (As you might have guessed, absolutely nothing about this in the
Hitachi manual either.)

On a separate note, I notice that, while the main filter for the unit
needs replacement every 6 months, the recommendation for the activated
charcoal odor filter for the oil mist filter unit is replacement monthly!
At 2 pumps X $45 each, monthly replacement sounds pretty spendy. (The
Hitachi service contract doesn't cover consumables used in maintenance.)
How long do you think I can actually go on replacing the charcoal filter?
(Obviously, if there's a noticeable oil smell, it's time.)

} I know what you mean about Hitachi manuals - I'm
} still sending in comments and corrections. But,
} if it helps, Hitachi is no worse than any of the
} other microscope companies, and better than some.

Yes, and the separate manual for the Varian pump doesn't discuss what one
would do with a vacuum system that was attached. In any event, I've
subsequently found out I can turn off the unit at air pressure if I need
to.

Peter

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North Carolina State University (NCSU) invites applications for a Transmission Electron Microscopist to oversee its transmission electron microscopy (TEM) facilities within the NCSU Analytical Instrumentation Facility. The facility supports a wide variety of physical sciences research and education programs at NCSU and within the Triangle Region. The Facility Manager will have primary responsibility for management and operation of:

FEI Titan G2 60-300 kV STEM/TEM (probe-corrected, monochromated, super-x),
200 kV JEOL 2010F S/TEM,
200 kV JEOL 2000FX.

Management will also include all auxiliary equipment, including energy dispersive x-ray spectrometers, electron energy loss spectrometers, CCD cameras and an extensive sample preparation laboratory. The Facility Manager will oversee the training, maintenance/calibration, educational, outreach and user-base development activities of the TEM facility and will assist in performing advanced microscopy analysis, when needed.

A Ph.D. in Materials, Physical Sciences or Engineering is required. The prospective candidate must have a strong background in the theory and practice of TEM microcharacterizaton with a minimum of 3 years post-graduate experience. The candidate should also have experience with vacuum technology, electronics, and computers. Strong interpersonal, communication and management skills are essential.

Interested candidates should submit a resume and cover letter at https://jobs.ncsu.edu/postings/10122. In addition, candidates should three people from whom letters of recommendation can be requested.

AA/EOE. In addition, NC State welcomes all persons without regard to sexual orientation. Persons with disabilities requiring accommodations in the application and interview process please call (919) 515-3148. Final candidates are subject to criminal & sex offender background checks. Some vacancies also require credit or motor vehicle checks. If highest degree is from an institution outside of the U.S., final candidates are required to have their degree verified at www.wes.org. Degree must be obtained prior to start date. NC State University participates in E-Verify. Federal law requires all employers to verify the identity and employment eligibility of all persons hired to work in the United States.

James LeBeau
Assistant Professor
Department of Materials Science & Engineering
North Carolina State University
tel: (919) 515.5049
jmlebeau-at-ncsu.edu

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Posted on Behalf of
Susan E. Anagnost, Ph.D.
Chair and Associate Professor
Dept. of Sustainable Construction Management and Engineering
Director, N.C. Brown Center for Ultrastructure Studies
SUNY College of Environmental Science and Forestry

The ad is here:
http://esf.interviewexchange.com/jobofferdetails.jsp?JOBID=32493

Title: Research Support Specialist
Department: Sustainable Construction Management
Salary Range: $40,000 Annual Salary
Duration: One year, renewable

Support the analysis of industrial samples for the N.C. Brown Center for
Ultrastructure Studies, SUNY-ESF; specifically to analyze air samples for
asbestos-type fibers using the phase contrast method (PCM; NIOSH 7400).
Other duties include sample preparation and training of personnel in the PCM
method of fiber analysis, routine maintenance of related equipment, and to
assist other N.C. Brown Center personnel with data compilation and
reporting.

Requirements:

Qualifications: A Bachelors' degree and completion of the NIOSH 582 course
or equivalent, and certification to analyze and train others to read air
samples for asbestos-type fibers with the NIOSH 7400 method. Minimum of one
year's experience in a NYSDOH-certified laboratory is required.

Tony

.........................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
Environmental, Health & Safety, Microscopy, & Forensic Engineering
pH2, LLC
5250 E US Highway 36, Suite 830
Avon, IN 46123
(317) 718-7020 Office
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Title-Subject: [Filtered] LaB6 filament shelf life?

Message: I have a 6 year old unused Denka LaB6 filament. It has been stored in a N2 cabinet the
whole time. It was opened 3 years ago, but not used. I have heard some say that these filaments
have a "shelf life" of only 3 years (which is why I didn't use it 3 years ago). Is this shelf life
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Title-Subject: [Filtered] Virtual Light Microscopy

Message: We are trying to put together a virtual version of one of the Microscope Fair stations. Our
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Title-Subject: [Filtered] UV, X-Ray, Infrared and Pigment Dating Tests for Oil Paintings

Message: I have an Ivan Ayvazovski Painting and I want to have UV, X-Ray, Infrared and Pigment
Dating Tests on my painting. Later on my art historian in Rusia will authenticate the painting with
the help of the test resulsts.

Please inform me list of companies ( laboratories)in USA performing UV, X-Ray, Infrared and Pigment
Dating Tests on Oil Paintings.

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Email: j.janssen-at-nki.nl
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Title-Subject: [Filtered] vitual light microscope

Message: There is a website concerning vitual light microscope with some nice images. Maybe you can
use that. http://virtual.itg.uiuc.edu/
Also the zebrafish atlas is a kind of vitual microscope;
http://zfatlas.psu.edu/view.php?s=85&z=1&c=6074,2818
Hope you can use them. I have no interest of any kind in these sites, Hans.

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Van,
Check Denka's site and see what they have for information. I'm more
familiar with Kimball and they have a wealth of information about LaB6.
See:

http://www.kimballphysics.com/cathodes-and-emitters/technology/technical-inf
ormation

In particular

LaB6-01 Technical Bulletin PDF - (1,653 KB) General Guidelines for
Operating LaB6 Cathodes

States that, " A rough, white, powder-like deposit on the cathode
itself indicates the cathode operating temperature is too
low (not even the oxide can be evaporated), and the
vacuum is excessively poor (too much oxide is being
formed). A gun which is vented to air (dry nitrogen is
preferred), without waiting for the gun structure to cool (a
few minutes is appropriate), causes the cathode to become
a dark blue to bluish silver color. Such a cathode can
often be reactivated by over-temperature operation at 2000
to 2100 K for 10 to 15 minutes; the surface layer,
presumably mostly LaB" evaporates away, exposing fresh
LaB6 underneath. As the emission reappears, it is
important to reduce back to a normal 1750 to 1850 K
operating temperature."

The idea of a shelf life may be related to the oxide buildup. I don't know
if Denka tips will survive the over-temp operation needed to clean the tip,
but I've never had any issues with (or statements about) LaB6 shelf life.
If the crystal is purple, I'd go with it. If it's blue, you've really not
got anything to lose in trying to recover it.

Also see:

LaB6-07 Technical Bulletin PDF - (335 KB) Recovery of Emission from ES-423E
LaB6 Cathodes Following a Vacuum Dump

This gives more detailed information about the things that can happen to a
tip and maybe still be recovered.

Disclaimer: I have no financial interest in Kimball, but prefer and
recommend to my customers, their LaB6 cathodes.

Ken Converse
Owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Title-Subject: [Filtered] LaB6 filament shelf life?

Message: I have a 6 year old unused Denka LaB6 filament. It has been stored
in a N2 cabinet the
whole time. It was opened 3 years ago, but not used. I have heard some say
that these filaments
have a "shelf life" of only 3 years (which is why I didn't use it 3 years
ago). Is this shelf life
real? Can I baby it into use with a slow outgassing process? Thanks to
anyone with feedback.

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Listers:

I just had a communication from Matt Belic (Matt Belic
{mimtnbiker-at-gmail.com} ) who has acquired possession of an RCA EMU-3
and an RCA EMU-4 electron microscopes, and who would like to find a
suitable way to dispose of them. He thinks they are in fairly good
condition, although they apparently have not been operated for some
time. If any of you know of anyone that would like to have one or
both of these instruments. please let Matt know. For one thing, both
instruments are now around 50 years old, and therefore might be
deemed to be antiques, so maybe some museum would like to have them.
If you would like more information about RCA's line of instruments go
to http://www.historic-glendale.net/microscope_history.htm,
--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

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2, 16 -- From: Wil Bigelow {bigelow-at-umich.edu}
2, 16 -- Subject: [Microscopy]RCA EMs available
2, 16 -- Cc: Matt Belic {mimtnbiker-at-gmail.com}
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On Jul 3, 2012, at 7:16 PM, bigelow-at-umich.edu wrote:

}
} I just had a communication from Matt Belic (Matt Belic
} {mimtnbiker-at-gmail.com} ) who has acquired possession of an RCA EMU-3
} and an RCA EMU-4 electron microscopes, and who would like to find a
} suitable way to dispose of them. He thinks they are in fairly good
} condition, although they apparently have not been operated for some
} time. If any of you know of anyone that would like to have one or
} both of these instruments. please let Matt know. For one thing, both
} instruments are now around 50 years old, and therefore might be
} deemed to be antiques, so maybe some museum would like to have them.
} If you would like more information about RCA's line of instruments go
} to http://www.historic-glendale.net/microscope_history.htm,
} --
} Wilbur C. Bigelow, Professor Emeritus
} Materials Sci. & Engr., Univ. of Michigan
} Ann Arbor, Michigan 48109-2136
} e-mail: bigelow-at-umich.edu;
} Fx:734-763-4788; Ph:734-975-0858
} Address mail to: 2911 Whittier Court
} Ann Arbor, MI 48104-6731

Dear Matt,
I suggest contacting local high schools. I know of several schools
in various parts of the country that have been interested in EMs in
the past.
Yours,
Bill

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Email: david.mitchell-at-sasol.com
Name: Dave Mitchell

Organization: Sasol

Title-Subject: [Filtered] Magnetic Fields

Message: Dear Listers
We are currently commissioning a new TEM lab and have been doing some preliminary field testing. The
lab EMI performance is good, but I'll do some detective work to try and improve it further. My query
concerns the rather unusual EMI frequency response we are getting. Normally the peak EMI field is
centred on 50Hz (60Hz in the US), due to the primary mains frequency. One then normally sees
harmonics thereof and these decrease in intensity in a roughly exponential manner with respect to
the mains peak. I see exactly that behaviour in both x and z directions but in my y direction the
100Hz signal is much more intense than the50Hz by a factor of 2. The origin is clearly external to
the lab, since I can electrically isolate the whole lab and this does not change. Does anyone have
any suggestions as to what sort of equipment/systems might be responsible for this 100Hz (double
mains) frequency?I'll be doing a full area survey next week, and so hopefully can track down the
source(s), but it would be good to know what candidate systems one should look at closely.
Thanks and regards,
Dave Mitchell

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Dave,
I would look for something that draws a fair amount of DC current. It's
going to have a full bridge rectifier and that will give you your doubled
frequency before it gets filtered.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Email: david.mitchell-at-sasol.com
Name: Dave Mitchell

Organization: Sasol

Title-Subject: [Filtered] Magnetic Fields

Message: Dear Listers
We are currently commissioning a new TEM lab and have been doing some
preliminary field testing. The
lab EMI performance is good, but I'll do some detective work to try and
improve it further. My query
concerns the rather unusual EMI frequency response we are getting. Normally
the peak EMI field is
centred on 50Hz (60Hz in the US), due to the primary mains frequency. One
then normally sees
harmonics thereof and these decrease in intensity in a roughly exponential
manner with respect to
the mains peak. I see exactly that behaviour in both x and z directions but
in my y direction the
100Hz signal is much more intense than the50Hz by a factor of 2. The origin
is clearly external to
the lab, since I can electrically isolate the whole lab and this does not
change. Does anyone have
any suggestions as to what sort of equipment/systems might be responsible
for this 100Hz (double
mains) frequency?I'll be doing a full area survey next week, and so
hopefully can track down the
source(s), but it would be good to know what candidate systems one should
look at closely.
Thanks and regards,
Dave Mitchell

Login Host: 196.34.16.68
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I have in the distant past plymerized LR White in the freezer with UV
light. My question is if this is possible/advisable at 4 degrees or even
at room temperature?

Aloha,
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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Hello, dear Tina,

as to my knowledge, LR White does polymerize with an exothermic reaction.
There are mentioned at least 4 possible procedures:
polymerized either chemically (by addition of a catalyst/accelerator)
by elevated temperatures (classical "polymerization oven" , or also by means of microwave treatment)
UV-irradiation (365 nm) in the cold (lowered temps 0�C- minus 20�)

Recommendations of most suppliers is to polymerize at least at 0�C down to -20�C, using UV light (30W, for approx. 24 h).

If you use(d) Chemical Accelerator (Cold Cure) this will end up with polymerization within at least 4-5 hours (-at- RT ) which to me seems to be a rather short and "speedy" polymerization process (and also has been reported to have poor infiltration properties). To achieve slower (and perhaps hence more equal) polymerization conditions one can place embedding capsules into ice. Polymerization by heat treatment (as usually done with epoxide resins) also is possible, the oven set to a temperature lower than 50�C for Gelatin capsules (for beem capsules suppliers tell us: 60-65�C).
(All polymerization in gelatin or beem capsules with tight closing by / firmly pressed parafilm-pieces over the cap (prevention of disturbing oxygen and moisture).

Hope it helps,
best wishes and good luck,

"Aloha" from Salzburg-Austria,

Wolfgang MUSS
EM-Lab
Univ.-Inst. Pathology
SALK-LKH(Gen. Hospital)
SALZBURG AUSTRIA

} -----Urspr�ngliche Nachricht-----
} Von: tina-at-pbrc.hawaii.edu [mailto:tina-at-pbrc.hawaii.edu]
} Gesendet: Freitag, 06. Juli 2012 20:43
} An: Mu� Wolfgang
} Betreff: [Microscopy] LR White UV embedding question
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} I have in the distant past polymerized LR White in the freezer with UV
} light.
} My question is if this is possible/advisable at 4 degrees or even at
} room temperature?
}
} Aloha,
} Tina
}
} ***********************************************************************
} *****
} * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu
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} * Biological Electron Microscope Facility * (808) 956-6251
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} * University of Hawaii at Manoa *
} http://www.pbrc.hawaii.edu/bemf*
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Dear Listers,

I inherited old Specimen Current Type 31 Meter/Amplifier, which I am
trying to put to good use, but unfortunately it refuses to power up
correctly.

I am wondering if someone may have its schematics and would be willing
to share it... Manual with specifications may also be helpful.

Thank you very much beforehand,
--
Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

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Name: Michael Cammer

Organization: NYU Langone Medical Center

Title-Subject: [Filtered] storing stained immunofluorescence in PFA?

Message: Today a student brought in a stained sample to image on the TIRF microscope. After she
stained the sample on Friday her supervisor told her that because the sample would not be imaged
until Monday, she should put it in fix, so she added the PFA to the sample. I asked whether she
misunderstood and her supervisor meant just the buffer (essentially PBS), but he really did mean the
2% PFA.

Does anyone else out there do this? Has anybody heard of this before?

Thank you.

Miohael.Cammer-at-med.nyu.edu

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If you store in PBS, you risk bacterial growth damaging the morphology. And both formaldehyde fixation and immunostaining is reversible so storage in fix seems like the appropriate thing to do. The only disadvantage is the need to do more rinsing before viewing.

Sent from my iPad

On Jul 9, 2012, at 8:24 AM, "microscopylistserver-noreply-at-microscopy.com" {microscopylistserver-noreply-at-microscopy.com} wrote:

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} Title-Subject: [Filtered] storing stained immunofluorescence in PFA?
}
} Message: Today a student brought in a stained sample to image on the
} TIRF microscope. After she stained the sample on Friday her
} supervisor told her that because the sample would not be imaged until
} Monday, she should put it in fix, so she added the PFA to the sample.
} I asked whether she misunderstood and her supervisor meant just the buffer (essentially PBS), but he really did mean the 2% PFA.
}
} Does anyone else out there do this? Has anybody heard of this before?
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Thanks to all who replied; I got 10 emails and 34 out-of-office replies.

I did not frame my question well enough, but it looks like everyone is
just about as confused as I am about certain things, and the tech sheets
about LR White from different vendors are inconsistent.

LR White now comes either with a *catalyst* already mixed in, or with the
catalyst (benzoyl peroxide) on the side that MUST be used for
polymerization.

The *accelerator* is an additional component that can be added for "Cold
Cure", which means polymerization at room temperature, which is strongly
exothermic (so Cold Cure is a bit of a misnomer).

LR White can be polymerized in an oven at } 65C if in polyethylene
capsules, or at 50-60C in gelatin capsules, and, according to come, as low
at 45C, but will not polymerize at 40C. I knew this; I have used 50C, but
my current project is sensitive to even this heat.

So LR White can be polymerized with UV. I have done this in the freezer
(-20C) in the past. My question was if it can be done at 4C (my
refrigerator is actually 6C). Several people wrote that it has polymerized
just fine at 4C, but now I realize I have no idea how exothermic this
reaction is, and if it is detrimental to antigen retention for
immunolabeling. That was really my question, but I just didn't day so!

The use of osmium tetroxide or even the presence of lots of metal (e.g.,
hemoglobin) can make the reaction so exothermic that stuff really gets
cooked. In addition, perhaps the optical density of osmicated tissue may
prevent the polymerization by UV.

Several people have the Pella UV cryo chamber and like it.

So I guess now the question is How exothermic is the polymerization at
each of the recommended temp/conditions? Could I have done it at 6C and
retained antigenicity?

Then there's the whole microwave polymerization thing, which I have also
done in the far past and didn't try here. I did polymerize in the freezer
with UV, but it took longer than I expected (48 hours), and I have no idea
how warm the tissue got.

Aloha,
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

==============================Original Headers==============================
12, 24 -- From tina-at-pbrc.hawaii.edu Mon Jul 9 14:50:17 2012
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Tina says " How exothermic is the polymerization at each of the recommended temp/conditions? Could I have done it at 6C and retained antigenicity?" My feeling is the reaction is pretty exothermic - I have seen polyethylene capsules deform from the heat of the reaction! I am 100% confident you can retain antigenicity at 6 C for antigen X but definitely not for antigen Y. Unfortunately, the only way to tell if you have antigen X or Y is to do the experiment. Armbruster et al. 1983 J. Histochem Cytochem 31:1380-1384 showed dehydration and embedding temperature influenced tubulin labeling at the EM level. See also my paper on a simple homemade device that uses a carbon dioxide gas cylinder to hold an aluminum block at between -20 and -45 C for processing and embedding tissues (Shoemaker et al., 2003 Microscopy Research and Technique 62:262-266). It is not as fancy as the Pella chamber but an inexpensive alternative.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: tina-at-pbrc.hawaii.edu [mailto:tina-at-pbrc.hawaii.edu]
Sent: Monday, July 09, 2012 2:51 PM
To: Phillips, Thomas E.

Thanks to all who replied; I got 10 emails and 34 out-of-office replies.

I did not frame my question well enough, but it looks like everyone is just about as confused as I am about certain things, and the tech sheets about LR White from different vendors are inconsistent.

LR White now comes either with a *catalyst* already mixed in, or with the catalyst (benzoyl peroxide) on the side that MUST be used for polymerization.

The *accelerator* is an additional component that can be added for "Cold Cure", which means polymerization at room temperature, which is strongly exothermic (so Cold Cure is a bit of a misnomer).

LR White can be polymerized in an oven at } 65C if in polyethylene capsules, or at 50-60C in gelatin capsules, and, according to come, as low at 45C, but will not polymerize at 40C. I knew this; I have used 50C, but my current project is sensitive to even this heat.

So LR White can be polymerized with UV. I have done this in the freezer
(-20C) in the past. My question was if it can be done at 4C (my refrigerator is actually 6C). Several people wrote that it has polymerized just fine at 4C, but now I realize I have no idea how exothermic this reaction is, and if it is detrimental to antigen retention for immunolabeling. That was really my question, but I just didn't day so!

The use of osmium tetroxide or even the presence of lots of metal (e.g.,
hemoglobin) can make the reaction so exothermic that stuff really gets cooked. In addition, perhaps the optical density of osmicated tissue may prevent the polymerization by UV.

Several people have the Pella UV cryo chamber and like it.

So I guess now the question is How exothermic is the polymerization at each of the recommended temp/conditions? Could I have done it at 6C and retained antigenicity?

Then there's the whole microwave polymerization thing, which I have also done in the far past and didn't try here. I did polymerize in the freezer with UV, but it took longer than I expected (48 hours), and I have no idea how warm the tissue got.

Aloha,
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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23, 29 -- From PhillipsT-at-missouri.edu Mon Jul 9 16:22:56 2012
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Hi,

from my experience with tissue cultures I can only say that too long
fixation can induce quite some background fluorescence, not sure how
much that applies to the sample in question. Also, PFA doesn't go along
well with all fluorophores. What I can recommend is using PBS with 0.1%
(w/v) NaN3 to avoid bacterial (or any other) growth. I kept fixed tissue
culture samples like that over a several weeks without any problem,
should also work for cells. Just keep in mind that NaN3 is quite toxic
and deal with it accordingly.

Kind regards,

Christian

On 09.07.2012 17:58, PhillipsT-at-missouri.edu wrote:
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} If you store in PBS, you risk bacterial growth damaging the morphology. And both formaldehyde fixation and immunostaining is reversible so storage in fix seems like the appropriate thing to do. The only disadvantage is the need to do more rinsing before viewing.
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} } Email: michael.cammer-at-med.nyu.edu
} } Name: Michael Cammer
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} } Organization: NYU Langone Medical Center
} }
} } Title-Subject: [Filtered] storing stained immunofluorescence in PFA?
} }
} } Message: Today a student brought in a stained sample to image on the
} } TIRF microscope. After she stained the sample on Friday her
} } supervisor told her that because the sample would not be imaged until
} } Monday, she should put it in fix, so she added the PFA to the sample.
} } I asked whether she misunderstood and her supervisor meant just the buffer (essentially PBS), but he really did mean the 2% PFA.
} }
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9, 24 -- From christian.liebig-at-medizin.uni-tuebingen.de Tue Jul 10 06:19:05 2012
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Hi, Tina:

Just out of curiosity, what made you think the antigen you are trying to
label is sensitive to heat? Thank you.

Hong

On 7/9/12 3:52 PM, "tina-at-pbrc.hawaii.edu" {tina-at-pbrc.hawaii.edu} wrote:

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Tina, it seems that your main concern is the maximum temperature during curing. That depends on many factors which seem to be getting jumbled together here.

If the reaction was not exothermic, there would be no issue. There would be no generation of heat to cause a problem. The only question would be the temperature conditions necessary to get the reaction to proceed.

However, given that the reaction is exothermic, the issue really becomes one of heat generation and heat transfer.
- A strongly exothermic reaction will definitely make the problem worse because more heat will be generated per unit of mass.
- A quick reaction will make the problem worse because that heat will be generated in a shorter time and will have less time to dissipate.
- More mass will make the matter worse because you have that many more units generating heat.
- Low heat conductivity will make the matter worse because once the heat is generated it will not be conducted away so well and will accumulate heating up the sample.

You can't do much about the first factor. It will be a function of the resin and its reaction. You should be able to control the latter three to try to keep the temperature within your bounds. However, I don't think that I want to analyze the problem to determine what the heat rise will be for the specific conditions. I'd rather just try to push it in the right direction.

You could use a case of an instantaneous reaction with no conduction to determine the worst case of temperature rise. Hopefully that is far worse than what you would see in practice.

You might be able to control the speed of the reaction by the combination of reactants. If the resin will still polymerize with less accelerator, maybe you can just give it more time.

You should also be able to control the speed of the reaction by the initial temperature of the reaction. I seem to recall the rule of thumb was a doubling of the rate for every 10C increase in temperature. Of course, an exothermic reaction will generate heat which will warm things up and speed things up which will lead to an even faster release of heat which will...

I have worked with small amounts of resin in my material applications. I could pot samples all day working with 1-inch molds. However, increasing the volume to 2-inch cubes led to situations where the resin heated to the point of smoking and nearly burning. We also don't think much of the exotherm involved with concrete curing. However, the mass of something like the Hoover Dam renders the exotherm a very real concern. I understand that was a major factor determining how fast the concrete could be placed - even with cooling lines.

Temperature sinks in conjunction with high heat conductivity can do a lot to limit the maximum temperature. It could well be that the limiting factor will be the resin itself. Much can be gained by keeping the mold thin so that the heat does not have so far to flow.

Like I said earlier, a full analysis would probably not be worthwhile. It brings back not-so-fun memories of partial differential equations from many years ago. Hopefully you can play with these factors and find some workable conditions.

Warren

-----Original Message-----
X-from: tina-at-pbrc.hawaii.edu [mailto:tina-at-pbrc.hawaii.edu]
Sent: Monday, July 09, 2012 2:51 PM
To: wesaia-at-iastate.edu

Thanks to all who replied; I got 10 emails and 34 out-of-office replies.

I did not frame my question well enough, but it looks like everyone is
just about as confused as I am about certain things, and the tech sheets
about LR White from different vendors are inconsistent.

LR White now comes either with a *catalyst* already mixed in, or with the
catalyst (benzoyl peroxide) on the side that MUST be used for
polymerization.

The *accelerator* is an additional component that can be added for "Cold
Cure", which means polymerization at room temperature, which is strongly
exothermic (so Cold Cure is a bit of a misnomer).

LR White can be polymerized in an oven at } 65C if in polyethylene
capsules, or at 50-60C in gelatin capsules, and, according to come, as low
at 45C, but will not polymerize at 40C. I knew this; I have used 50C, but
my current project is sensitive to even this heat.

So LR White can be polymerized with UV. I have done this in the freezer
(-20C) in the past. My question was if it can be done at 4C (my
refrigerator is actually 6C). Several people wrote that it has polymerized
just fine at 4C, but now I realize I have no idea how exothermic this
reaction is, and if it is detrimental to antigen retention for
immunolabeling. That was really my question, but I just didn't day so!

The use of osmium tetroxide or even the presence of lots of metal (e.g.,
hemoglobin) can make the reaction so exothermic that stuff really gets
cooked. In addition, perhaps the optical density of osmicated tissue may
prevent the polymerization by UV.

Several people have the Pella UV cryo chamber and like it.

So I guess now the question is How exothermic is the polymerization at
each of the recommended temp/conditions? Could I have done it at 6C and
retained antigenicity?

Then there's the whole microwave polymerization thing, which I have also
done in the far past and didn't try here. I did polymerize in the freezer
with UV, but it took longer than I expected (48 hours), and I have no idea
how warm the tissue got.

Aloha,
Tina

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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Hi, Hong-

I had the person who wants to do the TEM immunogold labeling first try her
antibody using fluorescence with some concentrations of glutaraldehyde in
paraformaldehyde (it can tolerate about 1% glut) and with heat (it does
not tolerate 50C).

X-from replies I've gotten it seems like any kind of polymerization of LR
White may be too exothermic! I will summarize more replies later...

Aloha,
Tina

} Just out of curiosity, what made you think the antigen you are trying to
} label is sensitive to heat? Thank you.
}
} Hong
}
} On 7/9/12 3:52 PM, "tina-at-pbrc.hawaii.edu" {tina-at-pbrc.hawaii.edu} wrote:
}
}
} } --------------------------------------------------------------------------
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} } LR White can be polymerized in an oven at } 65C if in polyethylene
} } capsules, or at 50-60C in gelatin capsules, and, according to come, as
} } low
} } at 45C, but will not polymerize at 40C. I knew this; I have used 50C, but
} } my current project is sensitive to even this heat.
} }
} } So LR White can be polymerized with UV. I have done this in the freezer
} } (-20C) in the past. My question was if it can be done at 4C (my
} } refrigerator is actually 6C). Several people wrote that it has
} } polymerized
} } just fine at 4C, but now I realize I have no idea how exothermic this
} } reaction is, and if it is detrimental to antigen retention for
} } immunolabeling. That was really my question, but I just didn't day so!
} }
} } Aloha,
} } Tina
}
}
} ________________________________
}
} This e-mail message (including any attachments) is for the sole use of
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} information. If the reader of this message is not the intended
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} prohibited.
}
} If you have received this message in error, please contact
} the sender by reply e-mail message and destroy all copies of the
} original message (including attachments).
}

****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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I just got my new issue of my favorite magazine (Make Magazine) and noticed a great article by Ben Krasnow on his home-built SEM from scratch. If anyone gets a chance, I highly recommend both the magazine in general, and this article.

He basically built an SEM from bare parts scrounged from other stuff. His column is exposed wire wrappings inside a bell jar.

--Justin A. Kraft

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Was it this guy?

http://www.popsci.com/diy/article/2011-03/video-diy-scanning-electron-microscope

Super impressive build. LOVE the spark plugs as electrical connections!

Tom Kaye

On 7/11/2012 3:16 PM, kraftpiano-at-gmail.com wrote:
} ----------------------------------------------------------------------------
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} I just got my new issue of my favorite magazine (Make Magazine) and noticed a great article by Ben Krasnow on his home-built SEM from scratch. If anyone gets a chance, I highly recommend both the magazine in general, and this article.
}
} He basically built an SEM from bare parts scrounged from other stuff. His column is exposed wire wrappings inside a bell jar.
}
} --Justin A. Kraft
}
}
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10, 18 -- From tom-at-TomKaye.com Wed Jul 11 17:59:45 2012
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Title-Subject: [Filtered] Electron Microscopy facilities in the northeast USA

Message: I am looking for any Electron Microscopy facilities that provide immunogold labeling
services around the New England and eastern midwest areas in the US. A biopharmaceautical company
in northern New Jersey has contacted us here in Georgia desiring to do some collaborative work with
an immunogold labeling project. We will take on this project only if there are no other EM
facilities closer to this company that they may collaborate their research. Thank you in advance
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Title-Subject: [Filtered] Microscopy and Microanalysis Impact Factor

Message: Thomson Reuter recently released 2011 Impact Factors and Microscopy and Microanalysis, the
Journal of the Microscopy Society of America, is again ranked as the top journal in microscopy
related disciplines with a 2011 IF of 3.01 and a 5-year IF of 3.38.

Microscopy and Microanalysis is an international journal that publishes original research reports
and topical reviews in all fields of microscopy, imaging and compositional analysis. Topics include
the development of instrumentation and techniques as well as applications associated with imaging
the microstructure and composition of samples in the biological and materials sciences.

In addition to publishing your research in a leading journal in the microscopy discipline
Microscopy and Microanalysis offers free printing of all color images, online publication of movies
and other supplementary data, no page charges, indexing in Pubmed/MEDLINE, and online publication
ahead of print.

Please consider MAM for publication of your research. Manuscripts can be submitted online at
http://mc.manuscriptcentral.com/mam. For further information please do not hesitate to contact me or
one of the other members of our editorial board.

Bob Price
Editor-in-Chief, Microscopy and Microanalysis
Bob.Price-at-uscmed.sc.edu

Bob Price
Tel: 803-216-3824
Admin Asst Tel: 803-216-3825
Dept Cell Biology and Anatomy
School of Medicine
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Here's the link to this guy's blog,
http://benkrasnow.blogspot.com/2011/02/first-photos-of-diy-scanning-electron.html.
He got tons of other interesting stuff, too.
Thanks to Tom and Justin for the post; it made my day.

On 7/11/2012 5:59 PM, tom-at-TomKaye.com wrote:
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} Was it this guy?
}
} http://www.popsci.com/diy/article/2011-03/video-diy-scanning-electron-microscope
}
} Super impressive build. LOVE the spark plugs as electrical connections!
}
} Tom Kaye
}
}
}
}
} On 7/11/2012 3:16 PM, kraftpiano-at-gmail.com wrote:
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} }
} } I just got my new issue of my favorite magazine (Make Magazine) and noticed a great article by Ben Krasnow on his home-built SEM from scratch. If anyone gets a chance, I highly recommend both the magazine in general, and this article.
} }
} } He basically built an SEM from bare parts scrounged from other stuff. His column is exposed wire wrappings inside a bell jar.
} }
} } --Justin A. Kraft
} }
} }
} }
} } ==============================Original Headers==============================
} } 5, 33 -- From kraftpiano-at-gmail.com Wed Jul 11 17:11:40 2012
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} } 5, 33 -- From: Justin Kraft {kraftpiano-at-gmail.com}
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} } 5, 33 -- Subject: Home built SEM.
} } 5, 33 -- Date: Wed, 11 Jul 2012 18:03:58 -0400
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} 10, 18 -- From tom-at-TomKaye.com Wed Jul 11 17:59:45 2012
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} 10, 18 -- From: Tom Kaye {tom-at-TomKaye.com}
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5, 28 -- From r-holdford-at-ti.com Wed Jul 11 23:44:15 2012
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Concerning safety: is it mentioned how X-Ray radiation produced by the e-beam is respected? Can the bell jar really give enough protection?
Marco

----------------------------------------
Marco M�ller
Platform Manager - Electron Microscopy

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11, 33 -- From mmoller-at-cicbiomagune.es Thu Jul 12 02:35:22 2012
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11, 33 -- Subject: RE: Home built SEM
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Hi all

As I've recently inherit an old Zeiss Photomicroscope II (~1969), I'm in
search for some manuals.

I've found these for the model I (~1960) et III (1975), but for mine,
I've only the general manual, which deals only with transmitted light.

So I'm looking for the manual ref G 41-655 for reflected light, and G
41-500 for polarised light.

If some would have a copy, to send me as a pdf file, I would be very
glad. No preference between french, english or german, but perhaps not
spanish or russian !

And by the way, if someone not too far from Strasbourg (France) has some
orphan spares for this microsocope, I'm interested too.

Thanks and best regards

Jacques

--

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Mat�riaux de Strasbourg
D�partement Surfaces et Interfaces
23, rue de Loess ; BP43
67034 Strasbourg CEDEX 2
France

Tel 00 33(0)3 88 10 71 01
Fax 00 33(0)3 88 10 72 48
E-mail : Jacques.Faerber-at-ipcms.unistra.fr

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I am trying to help an amateur who wants to section the down from
pigeon squabs.
Can anyone point me in the right methodological direction?

Thanks, Mike

--
Michael J. Herron, U of MN, Dept. of Entomology
herro001-at-umn.edu
612-624-3688 (office) 612-625-5299 (FAX)

==============================Original Headers==============================
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He is using low power LM. He wants to make some simple morph measurements.

On Fri, Jul 13, 2012 at 9:30 AM, Mu� Wolfgang {W.Muss-at-salk.at} wrote:
} Dear Mike,
}
} apologize if you find this not helpful�. but anybody will ask:
}
} grossing ¯o? (use knife)
}
} LM: Cryo?
}
} LM: Paraffin embedding/paraffinized sections/LM-staining
}
} TEM: resin embedding/resin sectioning and staining??
}
} Tem/SEM: Cryo-, CPD etc.,etc.
}
}
}
} Thank you for clarification (addendum sent best via Listserver again!),
}
}
}
} kind regards,
}
} Wolfgang MUSS PhD
}
} Salzburg-Austria
}
}
}
}
}
}
}
} Von: herro001-at-umn.edu [mailto:herro001-at-umn.edu]
} Gesendet: Donnerstag, 12. Juli 2012 21:17
} An: Mu� Wolfgang
} Betreff: [Microscopy] Down sectioning [ pigeon squabs = Daunen von
} Pinguin-Jungv�geln]
}
}
}
} ---------------------------------------------------------------------------
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}
}
} I am trying to help an amateur who wants to section the down from pigeon
} squabs.
}
}
}
} Can anyone point me in the right methodological direction?
}
}
}
} Thanks, Mike
}
}
}
} --
}
} Michael J. Herron, U of MN, Dept. of Entomology
}
} herro001-at-umn.edu
}
} 612-624-3688 (office) 612-625-5299 (FAX)
}
}
}
} ==============================Original Headers==============================
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}
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--
Michael J. Herron, U of MN, Dept. of Entomology
herro001-at-umn.edu
612-624-3688 (office) 612-625-5299 (FAX)

==============================Original Headers==============================
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List -eners,
By now, the thread is a bit frayed, but let me add a couple of stiches.

The issue is heat, more than temperature. Thus even though LR
white polymerization raises the temerature a lot, you can minimize
the amount of heat your sample gets by reducing the ambient
temperature, increasing the efficiency of heat exchange around your
sample and so on. It is definitely the case that antigens will suffer
a lot more heat if you polymerize LR white in a 50C box compared to a
4C box. Whether that lower amount of heat is still enough to gork
(scientific word meaning denature) your antigen of interst of course
depends on that antigen.

Besides lowering the temperature, it is also possible to
protect antigens. I found that adding DTT to mixures of butyl/methyl
methacrylate did not block polymerization but did prevent the sample
from turning brown during (also exothermic) polymerization and
helped a great many antigens survive. I have no idea what DTT would
do in LR white, but this is just an example of somthing that might
protect your antigen during the polymerization reaction.

As ever,
Tobias

--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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6, 21 -- To: tina-at-pbrc.hawaii.edu
6, 21 -- From: Tobias Baskin {baskin-at-bio.umass.edu}
6, 21 -- Subject: [Microscopy] Re: LR White UV embedding question
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Dear All:

If there is an evidence that heat is the problem in this case, the
labeling can be done before sample embedding. Thank you.

Hong

On 7/14/12 7:33 PM, "baskin-at-bio.umass.edu" {baskin-at-bio.umass.edu} wrote:

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Hi
I am looking to start a TEM lab in India. I heard that used TEMs are
available, sometimes free if one is willing to take it out of their
hands. Can someone give some information? I can pick it up in the USA.
I cannot afford a new one.
Thanks
Abdullah RN Thayyil

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Name: Michael Cammer

Organization: Skirball Institute of Biomolecular Medicine

Title-Subject: [Filtered] RE: [Microscopy] viaWWW:storing stained immunofluorescence in PFA?

Message: Thank you for the replies to my query about post-fixing fluorescent stained samples. The
replies that it may make the staining more stably bound and certainly prevents contamination may
make me a convert.

Regards,
Michael Cammer, Assistant Research Scientist
Skirball Institute of Biomolecular Medicine
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Dear Listers,

Can you advise on compressed air supply for FEI Vitrobot Mk3 plunger?
Should it be only the gas from cylinder or I can use a compressor? Do I
need to have any filters in line in case of compressor use?
Thank you in advance!

Sincerely,
Alex

--
Dr. Aleksandr Mironov MD, PhD
Senior Experimental Officer
D.1527, M.Smith Building
EM Core Facility, Faculty of Life Sciences
University of Manchester
Oxford Road
Manchester
M13 9PT
UK

Tel. +44-(0)161-275-5645
Fax. +44-(0)161-275-5171
E-mail: Aleksandr.Mironov-at-manchester.ac.uk
http://www.ls.manchester.ac.uk/research/facilities/electronmicroscopy/

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Hi everyone,

We've been tasked with looking at some porous powder metal compacts in
our TEM. The compacts are green (un-sintered) and so fall apart during
polishing. I tried crushing up a sample but the particles are too
thick to be electron transparent at 200 keV.

I'd like to try impregnating part of the compact with epoxy, followed
by sectioning and conventional polishing. Does anyone have any
suggestions on which epoxy to use and how best to accomplish this?

Thanks for your help!
--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering
Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

==============================Original Headers==============================
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Steven,

For vacuum infiltrating green powder metal parts I use Buehler EpoThin. Low viscosity, long time to polymerize (overnight at room temperature) so there is plenty of time to vacuum infiltrate. Mix it well, then mix it a little more - if it isn't mixed well it never hardens.

I use this for optical metallography (and sometimes the polished specimens wind up in the SEM too; beware of charging) but I'm not sure how well it would work for TEM.

Depending on the sample size your vacuum infiltration chamber can be pretty makeshift. I'll admit this here since y'all are such nice folks; I have a piece of silicon rubber sheet glued flat to an aluminum plate that forms the bottom of the vacuum chamber. The top is an inverted Pyrex funnel. Mechanical pump, rubber hose stepped down to neoprene fuel line from the auto parts store, that just fits the tubular part of the funnel. A little vacuum grease on the rim of the conical part, where it seals on the rubber base, and good-to-go.

If you push the neoprene line just a short way onto the tube, you can vary pumping rate by pinching / varying the angle - that is, you can throttle the vacuum and be gentle with the infiltration. Easy to make a mess if you get too enthusiastic; cleans up with acetone.

Have fun!

Regards,
Andrew Werner
Chief Metallurgist, Perforating
Schlumberger Reservoir Completions Center
14910 Airline Road, Rosharon, TX 77583-1590

"I Shoot the hippopotamus with bullets made of platinum,
because if I use leaden ones his hide is sure to flatten 'em"
The Bad Child's Book of Beasts - Hilaire Belloc

-----Original Message-----
X-from: spurgeon-at-drexel.edu [mailto:spurgeon-at-drexel.edu]
Sent: Tuesday, July 17, 2012 1:23 PM
To: Andrew Werner

Hi everyone,

We've been tasked with looking at some porous powder metal compacts in
our TEM. The compacts are green (un-sintered) and so fall apart during
polishing. I tried crushing up a sample but the particles are too
thick to be electron transparent at 200 keV.

I'd like to try impregnating part of the compact with epoxy, followed
by sectioning and conventional polishing. Does anyone have any
suggestions on which epoxy to use and how best to accomplish this?

Thanks for your help!
--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering
Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

==============================Original Headers==============================
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A number of years ago, Lucille Gianuzzi prepared some Mt. St. Helens ash for
TEM using the FIB. She basically welded the particles together with the Pt
and then thinned the sample. The technique should work for you too.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: spurgeon-at-drexel.edu [mailto:spurgeon-at-drexel.edu]
Sent: Tuesday, July 17, 2012 11:25 AM
To: s.walck-at-cox.net

Hi everyone,

We've been tasked with looking at some porous powder metal compacts in our
TEM. The compacts are green (un-sintered) and so fall apart during
polishing. I tried crushing up a sample but the particles are too thick to
be electron transparent at 200 keV.

I'd like to try impregnating part of the compact with epoxy, followed by
sectioning and conventional polishing. Does anyone have any suggestions on
which epoxy to use and how best to accomplish this?

Thanks for your help!
--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

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Email: watson-at-wi.mit.edu
Name: Nicki watson

Organization: whitehead institute

Title-Subject: [Filtered] formvar grids

Message: Dear List servers
I have been trying to make formvar sheets for two weeks with no success.
I normally have no trouble releasing thin formvar sheets from the glass slide, but for the past 2-3
weeks I am stuck. I realize we are in the dog days of summer and the humidity maybe part of the
problem but I am running out of my stock of grids!
I am using 0.3-0.5% Formvar in ethylene dichloride. Dipping the cleaned slides blotting 3-5 times
and drying in a jar with desiccant for 5mins to ½ hour.
I score the slide with a razor blade breath on it and slide it into clean water at a 45-degree
angle. My last attempt about 4 months ago resulted in over ten slides and a thousand grids. This
week nothing, the film will not release from the slide. I have tried every thing I know, old box of
slides, super clean slides, slides smeared with a bit of nose oil No luck. Any suggestions from
the old pros?
Thank you Nicki W

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Hello Nicki,
We had similar troubles some years ago when we had changed the supplier of the glass slides. We were not able to release a formvar film from these new slides.
Non of old tricks helped. Now, we are using shower gel for cleaning them and it works.

The procedure:
1. Put a drop of shower gel onto dry glass slide and carefully spread it over the glass slide surface with fingers (one minute procedure) .
2. Intensively wash the glass slide with warm tap water.
3. Put a drop of common kitchen detergent onto glass slide and spread it over the glass slide surface with fingers.
4. Intensively wash the glass slide with warm tap water (This step is important).
5. Briefly wash the glass slide with distilled water.
6. Wipe all water drops with filter paper and polish the glass slide with a piece of silk tissue or lens cleaning paper.
7. Immediately use for formvar film.

I hope it may help you.

Best regards Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

On Wednesday 18 of July 2012 07:02:19 you wrote:
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} Title-Subject: [Filtered] formvar grids
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} Message: Dear List servers
} I have been trying to make formvar sheets for two weeks with no success.
} I normally have no trouble releasing thin formvar sheets from the glass slide, but for the past 2-3
} weeks I am stuck. I realize we are in the dog days of summer and the humidity maybe part of the
} problem but I am running out of my stock of grids!
} I am using 0.3-0.5% Formvar in ethylene dichloride. Dipping the cleaned slides blotting 3-5 times
} and drying in a jar with desiccant for 5mins to � hour.
} I score the slide with a razor blade breath on it and slide it into clean water at a 45-degree
} angle. My last attempt about 4 months ago resulted in over ten slides and a thousand grids. This
} week nothing, the film will not release from the slide. I have tried every thing I know, old box of
} slides, super clean slides, slides smeared with a bit of nose oil� No luck. Any suggestions from
} the old pros?
} Thank you Nicki W
}
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Email: jmircheski-at-us.es
Name: Josif

Organization: University of Seville

Title-Subject: [Filtered] Viewing a diamond knife in SEM

Message: Dear Listers,

I am having some problems with my diamond knife, my
sections tend to have a lot of scratch marks. Probably the knife has a lot of scratches and the
useful surface of the edge is quite reduced. I am curious to see what is causing the scratches, is
it debris sticking to the edge or the edge itself is damaged. I'd like to check the knife in a SEM,
but I am not sure if the knife will be safe in the SEM conditions. Would the pressure or the beam
heat affect the diamond or the cement that is holding the diamond in any way?

Does anyone of you have any experience in seeing a diamond knife in SEM?

Thanks in advance,

Josif

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Hi Josif,

Thermal conductivity of diamond is *huge* and I can tell you from
practice that small diamond crystals survive FIB processing and SEM
observation just fine. I would not be too worried about damaging diamond
knife during regular SEM observation, at least not due to overheating by
the electron beam.

You have a different problem to worry about - diamond is dielectric and
will charge under the electron beam even more then the glass does.
Coating knife with metal or carbon to bleed the charge would ruin it, so
your best bet is to use environmental SEM and look at the knife in "high
pressure" mode under H2O atmosphere. You can try coating the knife with
washable conductive polymer for observation in high-vacuum SEM, but the
latter would hide tiny dents and scratches.

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
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Web: www.freudlabs.com

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} Title-Subject: [Filtered] Viewing a diamond knife in SEM
}
} Message: Dear Listers,
}
} I am having some problems with my diamond knife, my
} sections tend to have a lot of scratch marks. Probably the knife has a lot of scratches and the
} useful surface of the edge is quite reduced. I am curious to see what is causing the scratches, is
} it debris sticking to the edge or the edge itself is damaged. I'd like to check the knife in a SEM,
} but I am not sure if the knife will be safe in the SEM conditions. Would the pressure or the beam
} heat affect the diamond or the cement that is holding the diamond in any way?
}
} Does anyone of you have any experience in seeing a diamond knife in SEM?
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} Josif
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Years ago, I looked at a diamond studded dental drill. I coated the sample with gold for conductivity and to increase the secondary electron emission from the carbon based diamonds. It worked quite well.

And as a side affect, after seeing diamond studded drill bit, it increased my respect for dentist and my fear of the next visit (which is today.................)

Let us know how it works out.

Frank

-----Original Message-----
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Sent: Wednesday, July 18, 2012 8:20 AM
To: Frank Karl

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Email: jmircheski-at-us.es
Name: Josif

Organization: University of Seville

Title-Subject: [Filtered] Viewing a diamond knife in SEM

Message: Dear Listers,

I am having some problems with my diamond knife, my
sections tend to have a lot of scratch marks. Probably the knife has a lot of scratches and the
useful surface of the edge is quite reduced. I am curious to see what is causing the scratches, is
it debris sticking to the edge or the edge itself is damaged. I'd like to check the knife in a SEM,
but I am not sure if the knife will be safe in the SEM conditions. Would the pressure or the beam
heat affect the diamond or the cement that is holding the diamond in any way?

Does anyone of you have any experience in seeing a diamond knife in SEM?

Thanks in advance,

Josif
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This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Dear colleagues,
I just read this article and I cannot resist the urge to share it with you.
I find this article interesting because not only they reach very good resolutions even in the Z axis but also it is packed with methodological details and technical considerations.

http://www.biotechniques.com/BiotechniquesJournal/2012/July/High-resolution-three-dimensional-reconstruction-of-a-whole-yeast-cell-using----focused-ion-beam-scanning-electron-microscopy/biotechniques-332269.html?pageNum=1

regards,
Stephane

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A long time ago I put my wife's engagement ring in a high vac SEM without
coating. She was sure I was going to ruin it. I had no trouble at all
using 5kV and going to several thousand X to look at the edges of the
facets. I've heard that diamond is non-conductive, so perhaps it was the
accumulated dirt and grime on the stone that kept it from charging.

Guess I should try this again, then clean it and see what difference I get.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: frank_karl-at-ardl.com [mailto:frank_karl-at-ardl.com]
Sent: Wednesday, July 18, 2012 9:13 AM
To: kenconverse-at-qualityimages.biz

Years ago, I looked at a diamond studded dental drill. I coated the sample
with gold for conductivity and to increase the secondary electron emission
from the carbon based diamonds. It worked quite well.

And as a side affect, after seeing diamond studded drill bit, it increased
my respect for dentist and my fear of the next visit (which is
today.................)

Let us know how it works out.

Frank

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com
[mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: Wednesday, July 18, 2012 8:20 AM
To: Frank Karl

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Email: jmircheski-at-us.es
Name: Josif

Organization: University of Seville

Title-Subject: [Filtered] Viewing a diamond knife in SEM

Message: Dear Listers,

I am having some problems with my diamond knife, my
sections tend to have a lot of scratch marks. Probably the knife has a lot
of scratches and the
useful surface of the edge is quite reduced. I am curious to see what is
causing the scratches, is
it debris sticking to the edge or the edge itself is damaged. I'd like to
check the knife in a SEM,
but I am not sure if the knife will be safe in the SEM conditions. Would the
pressure or the beam
heat affect the diamond or the cement that is holding the diamond in any
way?

Does anyone of you have any experience in seeing a diamond knife in SEM?

Thanks in advance,

Josif
- Headers==============================

This email and any of its attachments may contain confidential information
intended only for the use of the addressee(s). If the reader of this email
is not the intended recipient or the employee or agent responsible for
delivering it to the intended recipient, you are hereby notified that any
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received this email in error, please notify us by return email at
info-at-ardl.com, permanently delete the email, and destroy any printouts. If
this email contains test data and/or draft reports, you are hereby notified
that only a signed original test report will contain official results, a
copy of which resides in the project folder located at ARDL, Inc. Thank you.
Akron Rubber Development Laboratory, Inc.

==============================Original Headers==============================
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Nicki,

Possible suggestion for "grid makers":
If one finds that grids are turning out well in the Spring, it is a good
idea to make twice as many as one had planned to make.
The humidity is usually much lower then so there are fewer holes in the
films, a big problem for me in both eastern Pennsylvania and Maryland, the
grids last for a half year and one will be using slides out of the same box.

X-from our in house supply store we can buy both standard and "Superfrost
Plus" slides. The latter have a cross (+) in the two end corners and are
made so that sections will adhere extra well. It is my experience that it
is impossible to get films off these slides, at least the one time that I
did not pay attention to the writing on the box that I had picked up.

The procedure that Oldrich has posted is one that I shall keep on file in
case I have problems with my standard glass slides in the future. I have
not tried his first step of using shower gel but using kitchen detergent
with washings has worked for me on old slides that I had found in the back
of a drawer.

Pat

Patricia Stranen Connelly
Research Assistant
NHLBI Electron Microscopy Core
National Institutes of Health
14 Service Road West
Bldg. 14E � Rm. 111B MSC 5570
Bethesda, MD 20892-5570
Phone 301-496-3491
FAX 301-402-0170
connellyps-at-mail.nih.gov {mailto:connellyps-at-mail.nih.gov}

Opinions and experiences related are those of Pat Connelly and do not
represent the NIH. This message is not confidential and can be freely shared
and reproduced.

============

X-from: {benada-at-biomed.cas.cz}
Reply-To: {benada-at-biomed.cas.cz}

Years ago I have observed an old diamond knife at 1 kV without coating. My main concern (rightfully or not) was heating under the beam and possible increase of adhesion of debris, so I used low mags (not higher than 1000). I have found out a lot of sections stuck to the edge. After soaking overnight in Alconox and cleaning with a foam stick, the knife worked again. It was rather difficult to remove all traces of Alconox; I had to wash it (soak) for two days with several changes of water. As an additional benefit I got excellent wetting of the edge of the knife.

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

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} Subject: [Microscopy] viaWWW:Viewing a diamond knife in SEM
}
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} Name: Josif
}
} Organization: University of Seville
}
} Title-Subject: [Filtered] Viewing a diamond knife in SEM
}
} Message: Dear Listers,
}
} I am having some problems with my diamond knife, my sections tend to have
} a lot of scratch marks. Probably the knife has a lot of scratches and the useful
} surface of the edge is quite reduced. I am curious to see what is causing the
} scratches, is it debris sticking to the edge or the edge itself is damaged. I'd
} like to check the knife in a SEM, but I am not sure if the knife will be safe in
} the SEM conditions. Would the pressure or the beam heat affect the
} diamond or the cement that is holding the diamond in any way?
}
} Does anyone of you have any experience in seeing a diamond knife in SEM?
}
} Thanks in advance,
}
} Josif
}
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On Jul 17, 2012, at 7:38 AM, Aleksandr.Mironov-at-manchester.ac.uk wrote:

} Dear Listers,
}
} Can you advise on compressed air supply for FEI Vitrobot Mk3 plunger?
} Should it be only the gas from cylinder or I can use a compressor?
} Do I
} need to have any filters in line in case of compressor use?
} Thank you in advance!
}
} Sincerely,
} Alex
}
} --
} Dr. Aleksandr Mironov MD, PhD
} Senior Experimental Officer
} D.1527, M.Smith Building
} EM Core Facility, Faculty of Life Sciences
} University of Manchester
} Oxford Road
} Manchester
} M13 9PT
} UK

Dear Alex,
We operated our Vitrobot using house compressed air. We incorporated
a regulator to achieve the recommended pressure, and the air was clean
enough that we didn't need a filter. However, I would definitely
recommend using a filter if there is any doubt as to the cleanliness
of the air supply. In particular, if the compressor you refer to is a
stand-alone apparatus located near the Vbot, I would definitely use a
filter.
Yours,
Bill

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Title-Subject: [Filtered] JEOL-6400 F

Message:
We are a R&D company based in the UK called Silicon CPV, part of the Akhter group of companies. We
have recently acquired a JEOL-6400 F scanning electron microscope from BP solar. However we now need
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July 30, 12:00pm (noon) Central Daylight Time (U.S.). For Photoshop
Extended versions CS3 (version 10) - CS6 (version 13). To sign up, go to:

http://www.imagingandanalysis.com/web072012ms.html

You will learn to post-process movie files and things like putting
movies side-by-side, adding arrows and other symbols, adding captions,
and so on. Here is a list of what will be shown:

- Open Image stacks as image sequences; or as a z- or time-series saved
in the uncompressed AVI format
- Maintain a log of what you did to movie files
- Open DICOM series as movies
- Add text, arrows, captions, etc. to movies
- Filter movies all at once: De-Interlace, Sharpen, etc.
- Place movies side-by-side; overlay movies to merge
- Adjust tonal range for optimizing to output devices
- Export completed movies for frame-by-frame "scientific" replay

Cheers,
Jerry

--
Jerry (Gerald) Sedgewick
Scientific Imaging & Image Analysis Consultant
Technical Writer / Regulatory Consultant
965 Cromwell Avenue
Saint Paul, MN 55114
651-788-2261
http://www.imagingandanalysis.com
http://www.quickphotoshop.com
Author: �Scientific Imaging with Photoshop: Methods, Measurement and Output�

==============================Original Headers==============================
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Absent an ion mill, I just pressed it well into the carbon double-stick when I finally got near a SEM.

The day after the explosion, my Mom and Dad found about an eighth to a quarter inch of dust on 'everything' outside their home in Sedona, Arizona - about 1300 miles SSE of the volcano - at about 4500 feet of elevation. Mom collected some and sent it in Gerber baby food jars to her two children in the East as part of our next Xmas boxes.

Phoenix(?????) - here I come - again!!! I remember well - deplaning (in the mid 1970's) directly to the tarmac and right after a mid-afternoon rain. I believe the tarmac dried before we had walked to the terminal. There was steam/mist everywhere - all of it rising. Every other climate I have visited since has been cool!!

Cheers,

Fred Monson

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
Schmucker Science Center South
West Chester University, West Chester, PA 19383
610-738-0437

Home Page: http://cmirt.wcupa.edu
Scheduler: http://cmirt.wcupa.edu/cgi-bin/ureserve_gold.pl

-----Original Message-----
X-from: S.Walck-at-cox.net [mailto:S.Walck-at-cox.net]
Sent: Tuesday, July 17, 2012 5:33 PM
To: Monson, Frederick

A number of years ago, Lucille Gianuzzi prepared some Mt. St. Helens ash for TEM using the FIB. She basically welded the particles together with the Pt and then thinned the sample. The technique should work for you too.

-Scott

Scott D. Walck, Ph.D.
5234 Sandalwood PL
Oceanside, CA 92056

S.Walck-at-cox.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(760) 758-4384 (Home)

-----Original Message-----
X-from: spurgeon-at-drexel.edu [mailto:spurgeon-at-drexel.edu]
Sent: Tuesday, July 17, 2012 11:25 AM
To: s.walck-at-cox.net

Hi everyone,

We've been tasked with looking at some porous powder metal compacts in our
TEM. The compacts are green (un-sintered) and so fall apart during
polishing. I tried crushing up a sample but the particles are too thick to
be electron transparent at 200 keV.

I'd like to try impregnating part of the compact with epoxy, followed by
sectioning and conventional polishing. Does anyone have any suggestions on
which epoxy to use and how best to accomplish this?

Thanks for your help!
--
Steven Spurgeon
Dynamic Characterization Group
Department of Materials Science & Engineering Drexel University

Office: Bossone 105
Email: spurgeon-at-drexel.edu
Web: http://www.materials.drexel.edu/dcg/

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Hello Wazz,

I probably can help you with answering all sort of questions you might have in connection with installation and operation of JEOL-6400F. The only thing you may not expect from me is to physically show up there and put together this SEM for you and train you guys how to operate, for the reason being, I am not by any means nearby you in UK. I am in USA. However, should you have someone there who could take care of the basic installation of 6400F but have questions or need additional guidance or get stuck somewhere and need help, I could provide this help remotely from here.

Zia ur Rahman
Johnson Space Center,
USA

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Email: wazz.mughal-at-akhter.co.uk
Name: Wazz Mughal

Organization: Silicon CPV

Title-Subject: [Filtered] JEOL-6400 F

Message:
We are a R&D company based in the UK called Silicon CPV, part of the Akhter group of companies. We have recently acquired a JEOL-6400 F scanning electron microscope from BP solar. However we now need to commission and operate it. Does anyone have any tips/pointers/experience in this?

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Dear All,

Thanks a lot for your suggestions on how to view a diamond knife in a SEM.
Unfortunately, we didn't have a ESEM at hand at the time, so we tried to
check it in low vacuum in a JEOL 6460LV with back-scattered electrons. The
resolution was quite low and I didn't see what I expected to see. The knife
does not show any visible signs of damage caused by the SEM, which was my
biggest worry and the reason I consulted the list.
I'll try the step Vlado suggested (hvala Vladimire), wash the knife in a
detergent and see if the scratches disappear.

Best,
Josif

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

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Email: jmircheski-at-us.es
Name: Josif

Organization: University of Seville

Title-Subject: [Filtered] Viewing a diamond knife in SEM

Message: Dear Listers,

I am having some problems with my diamond knife, my
sections tend to have a lot of scratch marks. Probably the knife has a lot
of scratches and the
useful surface of the edge is quite reduced. I am curious to see what is
causing the scratches, is
it debris sticking to the edge or the edge itself is damaged. I'd like to
check the knife in a SEM,
but I am not sure if the knife will be safe in the SEM conditions. Would the
pressure or the beam
heat affect the diamond or the cement that is holding the diamond in any
way?

Does anyone of you have any experience in seeing a diamond knife in SEM?

Thanks in advance,

Josif

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Email: craque-at-bergen.org
Name: Craig Queenan

Organization: Bergen County Technical Schools

Title-Subject: [Filtered] Education Outreach Session at M&M

Message: Hello All,

Microscopy and Microanalysis 2012 is less than a month away, and as a co-chair of X-90: Microscopy
in the Classroom Strategies for Education and Outreach, I wanted to reach out to the greater
microscopy community with information about the session.

This year, X-90 will have a roundtable format and feature three presentations on topics related to
implementing microscopy in K-12 classrooms and curricula, forms of academic and corporate
educational outreach, and engaging ways to introduce students to microscopy. Each presentation will
be followed by a discussion period to allow attendees to exchange stories of successes, programs of
interest, and to network with others involved in microscopy education or outreach events. Tabletop
SEM demonstrations will be available during the session break, as well as in-room refreshments.

The session is on Tuesday, July 31, from 8:30AM 12:00PM in room 229A of the Phoenix Convention
Center. It is our hope that members of the microscopy community will attend all or parts of the
session to not only support microscopy education, but also to share the knowledge and expertise that
they have acquired with years of microscopy experience. This knowledge is invaluable and critical
for improving methods of microscopy education at every level, and to hopefully leave a lasting
impact on the local educators attending in the greater Phoenix area.
More information about this session is available at the following link:

https://www.dropbox.com/s/w5vpzy087zzsnke/2012%20X90%20Program%20Publicity%20Flyer%20-%20P.pdf

I look forward to seeing you at X-90 in Phoenix! Thank you in advance for your participation.

Craig Queenan, C.E.M.T.
MSA Education Committee Education Outreach Subcommittee
2012 X-90 Session Co-chair

Bergen County Technical Schools
Nano-Structural Imaging Lab (NSIL)
Lab: (201) 343-6000 ext 3336
Office: (201) 343-6000 ext 4645
craque-at-bergen.org
www.bergen.org/NSIL

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Organization: Virginia Tech

Title-Subject: [Filtered] Service Center price list

Message: I would like to get an idea of the prices EM service centers charge for SEM and TEM sample
prep, and scope time per hour on the scopes. Also cost of critical point drying and sputter coating
runs.

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Hi Kathy,

You can see the price list of our SEM, TEM, Sample Prep facilities at the NUANCE Center, Northwestern University, including the Critical Point Dryer and Sputter coating runs in this page:

http://www.nuance.northwestern.edu/epic/Policy%20and%20Rate/index.html

Shuyou

_____________________________
Shuyou Li, Ph.D.
Electron Microscopist
NUANCE Center
Northwestern University
2220 Campus Drive, Cook Hall RM#2036
Evanston, IL 60208-3108, USA
Ph: (847) 491-6723
Fax: (847) 467-6573
Email: syli-at-northwestern.edu
http://www.nuance.northwestern.edu/
http://www.fom.northwestern.edu/

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Title-Subject: [Filtered] Service Center price list

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My son-in-law once made diamond knives; he now
makes diamond scalpels for eye surgery. He first
tried SEM for inspection, but he switched to a
200x "toy" QX-3 when they were first introduced.
He routinely photographs edges before shipping &
when returned for resharpening, which eliminates
lots of arguments about who did what to the edge.
One of the new handheld CCD scopes would provide
somewhat higher resolution, but the old QX-3
still works for Bill.

Caroline

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5, 21 -- Subject: Re: [Microscopy] : Viewing a diamond knife
5, 21 -- Cc: {microscopy-at-microscopy.com} ,
5, 21 -- Bill Ballard {bballard-at-diamondsurgicalproducts.com}
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I did some research in the early 1970's in collaboration with the DuPont
diamond knife group utilizing SEM and a special knife boat that allowed
removal of the diamond/steel shank for ease of viewing. The Cambridge
Stereoscan S4 did not have the capabilities of current instruments so I
was looking at secondary electrons with accelerating voltage of
10--15kV. The small radius at the edge tended to charge thus
compromising the imaging so I coated the knifes with gold:palladium in a
vacuum evaporator. I observed small lumps or particles near the cutting
edge of the diamond that were presumed to consist of epoxy resin
deposited during sectioning. I did not find a method capable of removing
the deposits without damaging the steel and epoxy mounting despite
considerable effort including ultrasound, detergents and possibly weak
acids (my memory is not clear about the details). These deposits clearly
correlated with knife lines seen in ultrathin sections examined with a
TEM. I was able to section with the coated knives and found that the
coating generally sloughed off and did not affect sectioning. My
conclusion was that these deposits were only removed during the vigorous
mechanical polishing used when resharpening the knives so I did not
publish these results.

Happy sectioning!

Larry

On 7/20/2012 9:30 AM, schooley-at-mcn.org wrote:
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} My son-in-law once made diamond knives; he now
} makes diamond scalpels for eye surgery. He first
} tried SEM for inspection, but he switched to a
} 200x "toy" QX-3 when they were first introduced.
} He routinely photographs edges before shipping &
} when returned for resharpening, which eliminates
} lots of arguments about who did what to the edge.
} One of the new handheld CCD scopes would provide
} somewhat higher resolution, but the old QX-3
} still works for Bill.
}
} Caroline
}
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} } Dear All,
} }
} } Thanks a lot for your suggestions on how to view a diamond knife in a SEM.
} } Unfortunately, we didn't have a ESEM at hand at the time, so we tried to
} } check it in low vacuum in a JEOL 6460LV with back-scattered electrons. The
} } resolution was quite low and I didn't see what I expected to see. The knife
} } does not show any visible signs of damage caused by the SEM, which was my
} } biggest worry and the reason I consulted the list.
} } I'll try the step Vlado suggested (hvala Vladimire), wash the knife in a
} } detergent and see if the scratches disappear.
} }
} } Best,
} } Josif
} }
} } Dr.�Josif�Mircheski

--
Larry Ackerman, Specialist
Electron Microscopy Lab
Manager, Diabetes & Endocrinology Research Center Microscopy Core
UCSF, Dept. of Anatomy, Rm S657
513 Parnassus Ave., Box 0452
San Francisco, CA 94143

larry.ackerman-at-ucsf.edu

415-999-4758

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Good Afternoon,

My Ultracut S Control Panel/ Box has died and cannot be repaired. The Ultracut S Microtome itself is in good working condition.

Does someone happen to have a spare or surplused Ultracut S Control Box that works, that they no longer need?

Sincerely,
Virginia Crocker

Virginia Tanner Crocker
Biologist
NIH, NINDS EM Facility
MSC4477
Bldg. 49, Room 3C58
49 Convent Drive
Bethesda, MD 20892
301-402-1568
301-480-1485 (fax)
crockerv-at-ninds.nih.gov (email)

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8, 27 -- Subject: Looking for a fully functional Ultracut S Control Box
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I have 2 problems with my Zeiss Axiophot (about 25 years old). The 100W halogen bulb burned out and I replaced it. I took the lamp housing off and projected it on the wall. I got the filament in focus with the mirror image just below it. it looked pretty good. I then removed the tube diffuser in the frame and remounted the lamp. When I removed the eyepiece and looked down the tube, I could see a portion of the filament in sharp focus crossing the field of view. What I saw was a single filament image - not the double image of the real and mirror image. I assume that is what I am supposed to see at this stage. I then replaced the diffuser. A brightfield photo doesn't look too bad but when I remove the slide (but don't change the focus) and just take a photo, the resulting "white" image is quite uneven when I look at it in Photoshop.� Any thoughts?

My other problem is that my microscope has a large part on top of the frame that housed the two 35 mm cameras and ports for the digital camera and 4x5 inch film camera. Not surprisingly, it has been awhile since I used any of the film cameras. When I remove the digital camera and look down on the microscope tube it was attached to, I can see a much of junk (dried water droplets?) on one of the glass surfaces. The dirty surface is probably a piece of glass or prism at the same level as the oculars. I can't reach it since there are prisms in the large camera housing that prevent access. I would like to remove the large camera housing but can't figure out how. No obvious screws.� Has anyone ever done this? I may be forced to pay a pro to clean this thing but would like to avoid it unless it is absolutely necessary.

Thanks for any suggestions on either of these two problems. Tom Phillips

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Tom,

I don't have an Axiophot, but we do have an old Reichert MeF3 (that is now unused, to my chagrin) with a 150W xenon lamp source. The particulars may be different between scopes but the requirements for lighting are the same: they need a bright but evenly diffused light source for imaging. Our light source has a movable condenser lens in front of the light source and a mirror behind the light source which can be adjusted in the left-right, up-down, and fore-aft directions. A general procedure for the setup of our light source is to: 1) Adjust the condenser lens adjusting knob to bring the burner/filament image into sharp focus. 2) With the side-to-side control for the reflecting mirror, adjust the burner/filament image so that it becomes clearly visible close (adjacent) to its mirror image. 3) With the up/down control for the reflecting mirror, set the burner/filament image and mirror image at the same height. 4) With the fore/aft control for the reflecting mirror, set the position in which the two burner/filament images are of the same size. 5) With the side-to-side control for the reflecting mirror, readjust until both images superimpose. 6) Now, adjust the condenser adjusting knob to achieve a uniform illumination (i.e., defocus the image to a uniform, evenly diffused light source). Hope this helps, Rick

-----Original Message-----
X-from: PhillipsT-at-missouri.edu [mailto:PhillipsT-at-missouri.edu]
Sent: Tuesday, July 24, 2012 4:05 PM
To: Richard A. Ross

I have 2 problems with my Zeiss Axiophot (about 25 years old). The 100W halogen bulb burned out and I replaced it. I took the lamp housing off and projected it on the wall. I got the filament in focus with the mirror image just below it. it looked pretty good. I then removed the tube diffuser in the frame and remounted the lamp. When I removed the eyepiece and looked down the tube, I could see a portion of the filament in sharp focus crossing the field of view. What I saw was a single filament image - not the double image of the real and mirror image. I assume that is what I am supposed to see at this stage. I then replaced the diffuser. A brightfield photo doesn't look too bad but when I remove the slide (but don't change the focus) and just take a photo, the resulting "white" image is quite uneven when I look at it in Photoshop.� Any thoughts?

My other problem is that my microscope has a large part on top of the frame that housed the two 35 mm cameras and ports for the digital camera and 4x5 inch film camera. Not surprisingly, it has been awhile since I used any of the film cameras. When I remove the digital camera and look down on the microscope tube it was attached to, I can see a much of junk (dried water droplets?) on one of the glass surfaces. The dirty surface is probably a piece of glass or prism at the same level as the oculars. I can't reach it since there are prisms in the large camera housing that prevent access. I would like to remove the large camera housing but can't figure out how. No obvious screws.� Has anyone ever done this? I may be forced to pay a pro to clean this thing but would like to avoid it unless it is absolutely necessary.

Thanks for any suggestions on either of these two problems. Tom Phillips

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Email: as-at-astonmet.com
Name: alan stone

Organization: Aston Metallurgical Services

Title-Subject: [Filtered] Scanner Aspect Ratio Problem

Message: Hi All,

I am inquiring for a friend who needs to replace an old flatbed scanner
and so far is finding that the new scanners do not maintain a 1:1 aspect
ratio. This is an issue as their lab deals with highly unique specimens.
Their data is researched and published, so it is crucial that
measurements made from the scanned files be accurate.

While we can script a PhotoShop action to correct for the distortion, it
would be better to find a scanner which maintains a proper aspect ratio.
If you know of such a scanner, please contact me off-line at
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Al Stone
ASTON Metallurgical Services

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Our probe & SEM holders are designed for 25mm and 30mm diameter polished
sections, and we had been using Buehler's "phenolic ring forms". However,
I've just learned that Buehler has dropped the metric versions and is
selling the imperial sizes only. I can probably adapt to 25.4mm but not
easily to 32mm. Struers has nothing equivalent � is anyone aware of
another possible equivalent?

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Genuinely, Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
cogito ergo ZzoooomM

This electronic communication is governed by the terms and conditions at
http://www.mun.ca/cc/policies/electronic_communications_disclaimer_2012.php

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6, 22 -- Subject: metric ring forms
6, 22 -- From: Michael Shaffer {mshaffer-at-mun.ca}
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You might be able to find Teflon tube with an inner diameter of 25 mm along
with Teflon rod with a 25 mm diameter and make your own forms.
Alternatively, a machine shop can easily make them cylinders out of solid
stock to any dimension.

Ron L

-----Original Message-----
X-from: mshaffer-at-mun.ca [mailto:mshaffer-at-mun.ca]
Sent: Friday, July 27, 2012 9:33 AM
To: lherault-at-bu.edu

Our probe & SEM holders are designed for 25mm and 30mm diameter polished
sections, and we had been using Buehler's "phenolic ring forms". However,
I've just learned that Buehler has dropped the metric versions and is
selling the imperial sizes only. I can probably adapt to 25.4mm but not
easily to 32mm. Struers has nothing equivalent { is anyone aware of another
possible equivalent?

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Genuinely, Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation Memorial University St. John's,
Newfoundland, Canada cogito ergo ZzoooomM

This electronic communication is governed by the terms and conditions at
http://www.mun.ca/cc/policies/electronic_communications_disclaimer_2012.php

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Listers,

Please DO NOT reply to me. I am forwarding this email for a client.
Please reply directly to Peter Shields at petesfrm-at-yahoo.com .
Not to the list, either.
Thanks!

} From: Peter Shields {petesfrm-at-yahoo.com}
}
} We are looking to obtain a STEM to fit these requirements:
}
} - HAADF detector.
}
} - Cs corrected system (any voltage) would be best, but a 200 kV
} system without CS corrector *might* suffice. Cs correction is
} strongly preferred.
}
} - Cold FEG, or Schottky source. Not tungsten or LaB6.
}
} - Ideally, a 200 or 300 kV Cs corrected STEM with Gatan HAADF
} detector and DF/BF detector.
}
} - Resolution of at least 0.2 nm
}
} - Alignments at 80kV or lower and ability to switch easily between
} TEM and STEM mode.
}
} We are interested in new, used in excellent condition, purchase or
} lease. Time share may be an option.
} Vendors welcome, please contact me directly.
}
} Peter F. Shields

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Thanks for the advice I have gotten so far. I made major progress on aligning my halogen bulb. My official Zeiss manual for this microscope gives what I know see as inappropriate advice on how to do this. The manual clearly shows an illustration in which the two coil images (real and mirror) are located adjacent to each other and not overlapping. In some ways this is similar to what I have always understood to do for a HBO (mercury) bulb since I understood putting HBO images on top of each other created too much heat. But someone sent me a link to a different Zeiss manual where they explicitly state the two halogen filament image should be superimposed on each other with the mirror image filling in the spaces between the filament coil of the real image. This makes more logical sense. Another major problem was that the X, Y, Z controls for the bulb and mirror are part of one frame that is housed inside an outer shell. It turns out that the screws that held the internal frame to the outer shell had loosened after 25 years and this meant the internal frame tilted downwards. When I tightened those screws I was able to center the filament nicely. When I take a digital image of a blank field (after setting up Kohler illumination) and then looked at the image in Photoshop, it is clear the center of the image is brighter than the edges. The pixel intensity was a bell-shaped curve that was a good 35 pixel intensities wide. I tried defocusing the lamp a little bit with the collector knob to spread the light more evenly but can't eliminate this pixel intensity spread. We looked at two Olympus microscopes in our core facility and see a similar surprisingly wide range of pixels. If I take a bright field image, it looks pretty good but I still find this unevenness disturbing. Does anyone know how homogenous of a field of illumination is reasonable to expect?

In regards to my problem with a dirty glass surface in my optical path, I succeeded in getting the camera housing (that has the ports for the two 35 mm cameras, 4x5 camera and port for my digital camera) off the top of my microscope frame. I can clearly see the dirty piece of glass is the one that links the bottom of the camera housing to the microscope frame. I can't figure out how to disassemble the camera box to get to the glass surface inside. I am reluctant to be too adventurous since I don't want to risk destroying my only camera port. I am going to contact Zeiss to see what they would charge to clean this or if they can tell me how to open the box.

Have a good weekend. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: PhillipsT-at-missouri.edu [mailto:PhillipsT-at-missouri.edu]
Sent: Tuesday, July 24, 2012 2:58 PM
To: Phillips, Thomas E.

I have 2 problems with my Zeiss Axiophot (about 25 years old). The 100W halogen bulb burned out and I replaced it. I took the lamp housing off and projected it on the wall. I got the filament in focus with the mirror image just below it. it looked pretty good. I then removed the tube diffuser in the frame and remounted the lamp. When I removed the eyepiece and looked down the tube, I could see a portion of the filament in sharp focus crossing the field of view. What I saw was a single filament image - not the double image of the real and mirror image. I assume that is what I am supposed to see at this stage. I then replaced the diffuser. A brightfield photo doesn't look too bad but when I remove the slide (but don't change the focus) and just take a photo, the resulting "white" image is quite uneven when I look at it in Photoshop.� Any thoughts?

My other problem is that my microscope has a large part on top of the frame that housed the two 35 mm cameras and ports for the digital camera and 4x5 inch film camera. Not surprisingly, it has been awhile since I used any of the film cameras. When I remove the digital camera and look down on the microscope tube it was attached to, I can see a much of junk (dried water droplets?) on one of the glass surfaces. The dirty surface is probably a piece of glass or prism at the same level as the oculars. I can't reach it since there are prisms in the large camera housing that prevent access. I would like to remove the large camera housing but can't figure out how. No obvious screws.� Has anyone ever done this? I may be forced to pay a pro to clean this thing but would like to avoid it unless it is absolutely necessary.

Thanks for any suggestions on either of these two problems. Tom Phillips

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Hi Folks,

I was asked by a local company if anyone knows anyone in the Minneapolis
area who does freelance SEM maintenance and could change a FEG tip on an
FEI XL30S SEM. They are trying to find cheaper options than going with FEI
service and would need it done in the next 2-3 weeks. It is a Schottky
type emitter
and they would probably buy the emitter unit from FEI as they are the only
suppliers so they were told. You can reply to me directly or to the
list and I will pass
on any recommendations.

Nick

--

Dr Nicholas Seaton

SEM Specialist
Characterization Facility

College of Science and Engineering

University of Minnesota

Shepherd Labs

100 Union Street SE

Minneapolis

MN, 55455

email: seato008-at-umn.edu

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16, 33 -- Subject: Re: SEM FEG tip change
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Nicholas,

Depending on the price that your friends would be charged for a new tip,
refurbishing old tip (i.e. replacing W needle with Zr coating) could be
a cost-effective option for the next tip replacement. Unfortunately
turnaround time for the refurbishment is 4 to 6 weeks, so even if they
have the old tip for refurbishment it would be impossible to get done in
3 weeks.

If you do not find anyone local in Minneapolis area - feel free to
contact me offline for quotation and customer references.

Good luck :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 7/27/2012 5:39 PM, seato008-at-umn.edu wrote:
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} Hi Folks,
}
} I was asked by a local company if anyone knows anyone in the Minneapolis
} area who does freelance SEM maintenance and could change a FEG tip on an
} FEI XL30S SEM. They are trying to find cheaper options than going with FEI
} service and would need it done in the next 2-3 weeks. It is a Schottky
} type emitter
} and they would probably buy the emitter unit from FEI as they are the only
} suppliers so they were told. You can reply to me directly or to the
} list and I will pass
} on any recommendations.
}
} Nick
}
}
} --
}
} Dr Nicholas Seaton
}
} SEM Specialist
} Characterization Facility
}
} College of Science and Engineering
}
} University of Minnesota
}
} Shepherd Labs
}
} 100 Union Street SE
}
} Minneapolis
}
} MN, 55455
}
}
}
} email: seato008-at-umn.edu
}
} ==============================Original Headers==============================
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} 16, 33 -- Subject: Re: SEM FEG tip change
} 16, 33 -- From: Nick Seaton {seato008-at-umn.edu}
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X-from: wadowska-at-upei.ca ()

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Email: wadowska-at-upei.ca
Name: Dorota Wadowska

Organization: Atlantic Veterinay College/UPEI

Title-Subject: [Filtered] H 7500 shutting down

Message: Hi,
I wonder if you could help me to find a cause of this problem. Our H
7500 shuts down 5 minutes after kV was turned on. It does not do that
when kV is off. When it shuts down I get a message: "stage unit port
error". After a minute or so it changes to comm. unit reset or at one
point HV unit reset. None of the controls on both panels work. After
clicking reset button the scope restarts the program, everything is
downloaded and checked. Sometimes even after pressing reset and
restarting the computer controls still dont work. I can't use digital
camera either (AMT). I get a message on a computer operating the camera
"check connection to ComPort1". Would any of you have any suggestion as
what might be the problem?
Thank you
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} --_000_F80E859D5560544CB8CDCD16C587B7E1A63C4CX10MBX11yuyaleedu_
} Content-Type: text/plain; charset="Windows-1252"
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} The Biological and Cryo Electron Microscopy Core Facility at the Yale
} Unive=
} rsity School Medicine is seeking a candidate for the research assistant
} pos=
} ition.
} The candidate will have opportunities to learn basic and advanced
} electron =
} microscopy. The job responsibilities will include the tissue/sample
} proces=
} sing procedures, sectioning and microscopic digital imaging. The
} candidate =
} needs demonstrate a sense of responsibility and attention to detail in
} expe=
} rimental work and record keeping. You will need the ability to
} concentrate =
} on the task at hand for a sustained period of time and remain highly
} motiva=
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} ate well with a diverse group of users as well as work collaboratively on
} a=
} team are essential. A bachelor degree in biology, chemistry or a related
} f=
} ield and 2-years of related work experience are required.
} For further information or to apply for the job please visit Yale
} Universit=
} y HR website
} at:http://www.yale.edu/hronline/careers/application/external/i=
} ndex.html, and enter STARS Requisition Number : 17734BR.
} I will be attending M&M meeting in Phoenix from July 29 =96 Aug. 2, and
} ava=
} ilable for interview or to answer questions.
}
} Best regards,
}
} Xinran Nick Liu, M.D. & Ph.D.
} Director of CCMI EM Core Facility
} Research Scientist in Cell Biology
} Yale University School of Medicine
} 333 Cedar Street, SHM I-E16A
} New Haven, CT 06520
} Office: 203.785.4050
} Lab: 203.785.5390
} http://medicine.yale.edu/ccmi/em
}
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} --|The Biological and Cryo Electron Microscopy Core Facility at the Yale
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} electron=
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} 12pt; "--|processing
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} s|--/span--||--span style=3D"line-height: 18px; font-size: 12pt;
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} span style=3D"line-height: 18px; font-size: 12pt; "--|demonstrate a sense
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} responsibility and attention to detail in
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} m=
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} --|For further information or to apply for the job please visit Yale
} Universi=
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} |--/span--||--span class=3D"Apple-style-span" style=3D"font-size: 16px;
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} font-si=
} ze: 16px; font-weight: normal; text-decoration: none; "--||--font
} class=3D"Appl=
} e-style-span" size=3D"4"--|Best regards,|--/font--||--/span--||--/div--|
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} ze: 16px; font-weight: normal; text-decoration: none; "--||--font
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} |--div style=3D"font-size: 14px; "--||--span style=3D"font-style: normal;
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} ze: 16px; font-weight: normal; text-decoration: none; "--||--font
} class=3D"Appl=
} e-style-span" size=3D"4"--|Xinran Nick Liu, M.D. &
} Ph.D.|--/font--||--/span--||--/d=
} iv--|
} |--div style=3D"font-family: Calibri, sans-serif; font-size: 14px;
} "--||--font si=
} ze=3D"4"--||--font class=3D"Apple-style-span" face=3D"'Monotype
} Corsiva'"--||--span=
} style=3D"font-style: normal; font-weight: normal; text-decoration: none;
} f=
} ont-size: 16px; font-family: Calibri, sans-serif; "--||--font
} class=3D"Apple-st=
} yle-span" face=3D"Monotype Corsiva"--|Director
} of |--/font--||--/span--||--/font--||--span
} class=3D"Apple-style-span" style=3D"fon=
} t-size: 16px; font-family: 'Monotype Corsiva'; "--|CCMI EM Core
} Facility|--/spa=
} n--||--/font--||--/div--|
} |--div style=3D"font-family: Calibri, sans-serif; font-size: 14px;
} "--||--span cl=
} ass=3D"Apple-style-span" style=3D"font-size: 16px; font-family: 'Monotype
} C=
} orsiva'; "--||--font size=3D"4"--|Research Scientist in Cell
} Biology|--/font--||--/span=
} --||--/div--|
} |--div style=3D"font-family: Calibri, sans-serif; font-size: 14px;
} "--||--font cl=
} ass=3D"Apple-style-span" face=3D"'Monotype Corsiva'"--||--span
} style=3D"font-st=
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} f=
} ont-family: Calibri, sans-serif; "--||--font class=3D"Apple-style-span"
} face=3D=
} "Monotype Corsiva" size=3D"4"--|Yale
} University School of Medicine|--/font--||--/span--||--/font--||--/div--|
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} "--||--font si=
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} class=3D"Ap=
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} style=3D"font-style: norm=
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} class=
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} Cors=
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} ==============================Original
} Headers==============================
} 8, 27 -- From xinran.liu-at-yale.edu Sat Jul 28 17:23:22 2012
} 8, 27 -- Received: from vm-emlprdomr-06.its.yale.edu
} (vm-emlprdomr-06.its.yale.edu [130.132.50.147])
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} 8, 27 -- Jul 2012 18:23:21 -0400
} 8, 27 -- From: "Liu, Xinran" |--xinran.liu-at-yale.edu--|
} 8, 27 -- To: "Microscopy-at-microscopy.com" |--Microscopy-at-microscopy.com--|
} 8, 27 -- Subject: EM research assistant opening at Yale University
} 8, 27 -- Thread-Topic: EM research assistant opening at Yale University
} 8, 27 -- Thread-Index: AQHNbQ+Y8HmHgJC/ikyebwFh6eS5Dg==
} 8, 27 -- Date: Sat, 28 Jul 2012 22:23:20 +0000
} 8, 27 -- Message-ID:
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==============================Original Headers==============================
8, 30 -- From xinran.liu-at-yale.edu Sat Jul 28 17:26:36 2012
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8, 30 -- From: "Liu, Xinran" {xinran.liu-at-yale.edu}
8, 30 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
8, 30 -- Subject: FW: EM research assistant opening at Yale University
8, 30 -- Thread-Topic: EM research assistant opening at Yale University
8, 30 -- Thread-Index: AQHNbQ+Z0vVLSVX1WkyHLdj2pVAM6Zc/RfaA
8, 30 -- Date: Sat, 28 Jul 2012 22:26:34 +0000
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The Biological and Cryo Electron Microscopy Core Facility at the Yale
University School Medicine is seeking a candidate for the research
assistant position.
The candidate will have opportunities to learn basic and advanced electron
microscopy. The job responsibilities will include the tissue/sample
processing procedures, sectioning and microscopic digital imaging. The
candidate needs demonstrate a sense of responsibility and attention to
detail in experimental work and record keeping. You will need the ability
to concentrate on the task at hand for a sustained period of time and
remain highly motivated. Effective interpersonal (written and oral)
skills, ability to communicate well with a diverse group of users as well
as work collaboratively on a team are essential. A bachelor degree in
biology, chemistry or a related field and 2-years of related work
experience are required.

For further information or to apply for the job please visit Yale
University HR website
at:http://www.yale.edu/hronline/careers/application/external/index.html,
and enter STARS Requisition Number : 17734BR.

I will be attending M&M meeting in Phoenix from July 29 � Aug. 2, and
available for interview or to answer questions.

Xinran Nick Liu, M.D. & Ph.D.
Director of CCMI EM Core Facility
Research Scientist in Cell Biology
Yale University School of Medicine
333 Cedar Street, SHM I-E16A
New Haven, CT 06520
Office: 203.785.4050
Lab: 203.785.5390
http://medicine.yale.edu/ccmi/em

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X-from: m.aqel-at-mail.utoronto.ca ()

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Email: m.aqel-at-mail.utoronto.ca
Name: M.Aqel

Organization: Engineer

Title-Subject: [Filtered] Oxford X-ray analysis

Message: Hi,
I have a problem with the OXFORD INCAx-sight with ISIS link (Model
Number 23356.001). Whenever I want to run the ISIS application and I
want to run the x-ray analysis the following message appears:
-----------------------------------------
"This application will continue but with reduced functionality
Reason:
Application requires ISIS hardware
This application is entering demonstration mode since communications
with the ISIS system has not been established."
-----------------------------------------------

I run the Translink validation and the results were:
Card Test Passed
Cable Loopback Failed

the OXFORD system installed on a Hitachi SEM and the operating system in
the computer is Windows 3.11 for workgroup

any one has ever encounter such a problem or know how to fix please advise

Thank you
M.AQEL
Civil Engineering Department
University of Toronto

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Thanks all to who responded � but a quick clarification �

Those who did respond provided examples of tubing that I might simply
slice to suit. However, I'd question the materials mentioned (e.g.,
Teflon, Silastic) with regard to their adherence to epoxy(?) That is,
epoxy adheres to Buehler's phenolic ring forms, which makes them part of
the section after curing � teflon on the other hand would not adhere.

Still, your suggestions leave me with plenty of materials to explore, but
the questions would then become relative to properties like out-gassing,
etc. I use these ring forms for mounting the smallest of particles. That
is, the particles go on ultra-flat double-stick tape and the rings are
used to pour epoxy into. (Altho the possibility also exist to use
Silastic tubing with a 25mm ID � instead of 25mm OD phenolic forms � but
25mm OD forms still are preferred)

~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Genuinely, Michael Shaffer
SEM-MLA Research Coordinator
Bruneau Centre for Research & Innovation
Memorial University
St. John's, Newfoundland, Canada
cogito ergo ZzoooomM

On 12-07-27 11:30 AM, "lherault-at-bu.edu" {lherault-at-bu.edu} wrote:

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Interesting. We use Buehler blue plastic rings and coat the inner part
with a bit of petroleum jelly so that the epoxy can be removed. If you are
looking for a one piece result, you could even check out plastic plumbing
pipe to see if anything has the dimension you require. Then you'd only have
to cut them to length and dress the end, easily accomplished with a mini
lathe.

Ron L

-----Original Message-----
X-from: Michael Shaffer [mailto:mshaffer-at-mun.ca]
Sent: Monday, July 30, 2012 9:52 AM
To: MSA Microscopy list
Cc: lherault-at-bu.edu

I've gone both ways with my forms, I used to use the bakelite rings but have recently been favoring the blue, two-part, reusable molds.

I think the plastic pipe might be used as described below. It might also require dressing the outside of the pipe to get the size you want. The PVC that I am familiar with is quite irregular. But dressing the outside shouldn't be that hard.

However, if you have a machine shop available, it might be worth converting over to English sizes to be able to simply use the available rings.

Good luck with the hunt and let us know if you find a source for the metric forms.

Warren

-----Original Message-----
X-from: lherault-at-bu.edu [mailto:lherault-at-bu.edu]
Sent: Monday, July 30, 2012 7:01 AM
To: wesaia-at-iastate.edu

Interesting. We use Buehler blue plastic rings and coat the inner part
with a bit of petroleum jelly so that the epoxy can be removed. If you are looking for a one piece result, you could even check out plastic plumbing pipe to see if anything has the dimension you require. Then you'd only have to cut them to length and dress the end, easily accomplished with a mini lathe.

Ron L

-----Original Message-----
X-from: Michael Shaffer [mailto:mshaffer-at-mun.ca]
Sent: Monday, July 30, 2012 9:52 AM
To: MSA Microscopy list
Cc: lherault-at-bu.edu

Dear colleagues

As a reminder, this is final two days for abstract submission for the 3d
International workshop on "Imaging for Energy" applications, to be held
in conjunction with the CNMS User meeting. The information of the
workshop can be found at

http://cnms.ornl.gov/workshops/2012/SPM_EnergyWorkshop2012.pdf

Looking forward to seeing you in Oak Ridge

Sergei V. Kalinin

*REMINDER**� **Abstract deadline is Wednesday, August 1***

**

*CNMS User Meeting*

*Call for Abstracts for Oral and Poster Presentations
Abstract Deadline: August 1, 2012*(Click here
{https://register.ornl.gov/2012/cnms/poster_abstract.cfm} to submit an
abstract)
* Meeting Date: September 14, 2012
Poster Session and Reception: Thursday evening, September 13*

**

Researchers planning to attend the CNMS User Meeting are encouraged to
submit abstracts describing recent research results to be considered for
presentation in oral or poster sessions during the meeting. CNMS users
are especially encouraged to submit abstracts that include results from
their user projects but forefront research across the spectrum of
nanoscale materials science spanned by CNMS research capabilities and
user interests will be welcome. The CNMS User Executive Committee will
select a limited number of contributed abstracts for oral presentation
on Friday, September 14, to complement the invited talks. Additional
abstracts will be allocated space in the poster exhibit that will be on
display from Thursday afternoon through the day on Friday.

Abstracts may be prepared as either PDF or MS Word files and uploaded
using the online abstract submission form
{https://register.ornl.gov/2012/cnms/poster_abstract.cfm} . Abstracts
must not exceed one page in length, including figures, with one-inch
margins and minimum 11-pt font size.

* https://register.ornl.gov/2012/cnms/poster_abstract.cfm

*

*The deadline for abstract submission is August 1, 2012*. Submitting
authors will be notified of acceptance by August 15.

**

*REMINDER: Meeting registration deadline is August 31, 2012.
*A single registration fee covers all events including tutorials and
workshops on September 11-13. See meeting website
{http://www.cnms.ornl.gov/workshops/2012/announcement.shtm} for more
details. Click here
{https://register.ornl.gov/2012/cnms/poster_abstract.cfm} to register.

http://www.cnms.ornl.gov/workshops/2012/announcement.shtm

*
*

*https://register.ornl.gov/2012/cnms/poster_abstract.cfm
*

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My apologies, but the webinar service wouldn't start the webinar re:
Working with Movies and Image Stacks in Photoshop CS3 - CS5 at noon
today, and for which almost 200 people signed up. I will announce this
again after I sign up with a reliable webinar service.

Again, my apologies.

Jerry Sedgewick

--
Jerry (Gerald) Sedgewick
Scientific Imaging & Image Analysis Consultant
Technical Writer / Regulatory Consultant
965 Cromwell Avenue
Saint Paul, MN 55114
651-788-2261
http://www.imagingandanalysis.com
http://www.quickphotoshop.com
Author: �Scientific Imaging with Photoshop: Methods, Measurement and Output�

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5, 35 -- *CANCELED*
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Email: zerfasp-at-ors.od.nih.gov
Name: Patricia Zerfas

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Title-Subject: [Filtered] EM film and paper

Message: I have nine boxes of Ilford multigrade IV RC deluxe paper
glossy and one box of Kodak 4489 EM film (Qty 250) that I no longer can
use. I could ship it to your address if you provide an account number
and address. I am located in the USA. These items were kept in the
freezer since being ordered about three years ago.

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Email: sjlaugero-at-hotmail.com
Name: Silvio Laugero

Organization: Microscopy Laboratory Applied to Molecular Studies

Title-Subject: [Filtered] Questions

Message: Good morning: Our institution has a Hitachi HHS-2R SEM, which
is in operation. I wonder through this medium if anyone has parts of
this team and especially some EDX Analisys Device, original or that can
be adapted.
Thanks and best regards

Silvio Laugero.
Universidad Nacional de Entre Rios.
Argentina

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Our lab had just started doing some cryo-TEM of protein and virus suspensions. We have an investigator who is interested in doing single particle reconstruction of a purified protein complex. Does anyone know of any workshops, classes, or facilities that would provide training on single particle reconstruction?

Thanks,

Shannon

Delaware Biotechnology Institute
BioImaging Center
15 Innovation Way Suite 117
Newark, DE 19711

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Dear All,

Our old ANS EDS system mounted on our SEM (CAMSCAN SERIES II) is now
defunct and we are exploring the possibility of replacing it with a system
that won't cost us an arm and a leg (maybe just an arm or a leg. Well
better just a pinkie).

Does anybody have any suggestion regarding manufacturers or sources for
used/refurbished systems? The system will have to have a 45 degree take
off angle.

Thanks.

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
----------------------

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I have a visitor who wants to perform electron microprobe analysis on geologic thin sections mounted with canada balsam. I have lots of experience with epoxy mounted samples, but have never worked with canada balsam. Is canada balsam stable in high vacuum and under the heat of an electron beam? I'd appreciate comments from anyone who has experience working with canada balsam mounted samples.
--
Chris Fleisher
Electron Microprobe Lab Coordinator
Department of Geology
University of Georgia
Athens, Ga. 30602
Ph: (706) 542-2419 E-mail: fleisher-at-gly.uga.edu

==============================Original Headers==============================
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Hi all,

I'm looking for some assistance in troubleshooting/operating a GW
Electronics BSE detector system 47. It is connected to a Hitachi 4700
and has worked in the past, but there aren't any current users that
know how to operate the detector, and I can't get much beyond noise
out of it. The internet has been of little help so far, and it appears
as though the company is fading out of existence.

I'm wondering if anyone has a manual or other reference material for
this system, or is willing to help out with some basic operation and
troubleshooting tips?

Thanks in advance,

Edward Swanson
PhD Candidate
Columbia University
ejs2163-at-columbia.edu

==============================Original Headers==============================
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Dear All,
I am looking for a working PCI interface card for the O-SIS MegaViewII TEM camera.
See a small image of the card here:
www.electronmicroscopy.info/mv_12.jpg

I do not know any specifications of the card but it is not available any more from the manufacturer.

Thanks,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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Hi Edward

Before you worry too much about the detector let's check that it is working,
try these steps -

1. Move to a 15mm working distance as the ideal distance is said to be
12mm from the detector.
2. Turn the gain to its highest level (by memory it has three levels)
3. Crank up the spot size/probe current as high as possible
4. Make sure all of the segment switches are on positive
5. Slow the scan

If the above does not work you have a real problem!

Good luck

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: ejs2163-at-columbia.edu [mailto:ejs2163-at-columbia.edu]
Sent: 04 August 2012 19:02
To: protrain-at-emcourses.com

Hi all,

I'm looking for some assistance in troubleshooting/operating a GW
Electronics BSE detector system 47. It is connected to a Hitachi 4700 and
has worked in the past, but there aren't any current users that know how to
operate the detector, and I can't get much beyond noise out of it. The
internet has been of little help so far, and it appears as though the
company is fading out of existence.

I'm wondering if anyone has a manual or other reference material for this
system, or is willing to help out with some basic operation and
troubleshooting tips?

Thanks in advance,

Edward Swanson
PhD Candidate
Columbia University
ejs2163-at-columbia.edu

==============================Original Headers==============================
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Hello,

In early 2012, iLab Solutions conducted its second annual Core
Facility Benchmarking Study. In total, there were 208 individual
responses which represent 40 different core types from over 75
institutions from around the world. The analyzed data reveals a number
of strong patterns in core management and operations as well as
explores the biggest challenges core managers face today.

If you would like to view the full report, it can be downloaded at
http://www.ilabsolutions.com/news/2012-benchmarking-study-results/.

Kind regards,

Alicia Cravens
Marketing Manager
iLab Solutions, LLC
www.ilabsolutions.com

Meet iLab:
AIRI, Philadelphia (September 29-October 3)
DGFZ, Bonn (October 10-12)
AACI, Chicago (October 14-16)
MWACD, St. Louis (October 25-27)
WACD, Portland (November 1-2)

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I just wanted to thank you all for the replies and the info you supplied.
Now we have a base for starting our search for the new system. Hopefully
we'll find the money too :-)

Luca

-------------------------
Luca Fedele PhD
Department of Geosciences
4044 Derring Hall
Virginia Tech
Blacksburg, VA, 24061
lfedele-at-vt.edu
www.lucafedele.eu
----------------------

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5, 34 -- Subject: EDS for SEM - THANKS
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Dear Microscopists,
Hope folks had a great time at the MSA meeting and
are not too burried upon return.

This summer, I had the pleasure of doing some SEM in Japan.
The lab I visited prepared therir biological samples by freeze drying
out of tert-butyl alcohol (instead of critical point drying out of
CO2). This method (see Inoue and Osatake 1988 Arch Hitol Cytol. 51:
53-59) is widespread in Japan, but does not seem to be particularly
widely used in USA. Am I mistaken about that?

I am would like to hear from people who have tried freeze
drying out of tert-butyl alcohol for preparing samples for SEM. Good
experiences? bad ones? Comments??

Many thanks,
Tobias
--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

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I just wanted to follow up on the problem I had with a dirty internal lens on my camera housing unit. My local Zeiss rep John Dirnberger stopped by and showed me how to open up the housing for the unit that connected the camera to the microscope. I had been too timid and hadn't removed the slider arm and that is what was preventing my being able to remove the bottom cover. No problem once that arm was unscrewed. As we were opening the case, he raised the possibility that it was a coating coming off the lens but thankfully it was only an oily residue. I have no idea of how it got there. The bracket for glass lens mounted on top of this one had a grease layer holding it in place and maybe it just evaporated and re-deposited there over the millennia that I have been doing microscopy. We were able to clean it off. But the gods weren't done messing with me. The hard drive of the PC that drives the system's camera and stores its images failed almost immediately afterward. Fortunately my incredible IT department backs this system up automatically every 2 days. I guess if microscopy was easy, everybody would do it....

Thanks to all who sent advice and tips. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Listers,
There is an opening for a Director of the electron microscopy core facility at the University of Texas Southwestern Medical Center at Dallas.

Full information and contact details are available at http://www4.utsouthwestern.edu/mcif/director/coredirector.pdf

The Core's website is http://www4.utsouthwestern.edu/mcif

Regards

Chris

Christopher J Gilpin Ph.D.
Assistant Professor
Director, Molecular and Cellular Imaging Facility
K1.A04
UT Southwestern Medical Center at Dallas
5323 Harry Hines Boulevard
Dallas, TX 75390-9039
Phone +1 214 648 2827

________________________________

UT Southwestern Medical Center
The future of medicine, today.

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***************************************************************************************
Forwarded from "Ask a Microscopist"
Please remember that the person asking the question is likely
not a member the listserver, and
**any reply should go directly to the poster**
as well as to the list.
Using the "reply" function in your email does *not* send your answer
to the person asking the question.
Please copy their email address from their question.
****************************************************************************************
} realname - Susan C. Van Horn
} Email - susan.vanhorn-at-stonybrook.edu
} ORGANIZATION - SUNY -at- Stony Brook
} EDUCATION - Graduate College
} LOCATION - StonyBrook, New York, USA
} SUBJECT_OF_QUESTION - Agarose embedding of virus particles
} QUESTION - Does anyone have a protocol for embedding a virus pellet
} in agarose???........and is it better to fix before or after
} embedding the pellet in the agarose??? I have been reading if you
} fix after the pellet is in the agarose the glutaraldehyde cross-link
} is so strong that other chemicals may not penetrate.......
} thanks,
} sue

--

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Short answer - Fix before AND after.

Fix, spin into a pellet, refix, get into agarose, fix some more. Cut into
smaller cube for osmium tetroxide post-fixation. You should be OK if the
agarose bit is no larger than 1mm in one dimension so the chemicals can
diffuse in. Dehydrate and embed as usual!

Aloha,
Tina

} } SUBJECT_OF_QUESTION - Agarose embedding of virus particles
} } QUESTION - Does anyone have a protocol for embedding a virus pellet
} } in agarose???........and is it better to fix before or after
} } embedding the pellet in the agarose??? I have been reading if you
} } fix after the pellet is in the agarose the glutaraldehyde cross-link
} } is so strong that other chemicals may not penetrate.......
} } thanks,
} } sue
}
}
} --
}
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****************************************************************************
* Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 *
* University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
****************************************************************************

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Working with Image Stacks and Movies webinar is now re-scheduled with a
different webinar agency (so it works this time):

August 20, 12:00pm (noon) until 1:30pm Central Daylight Time (U.S.).
For Photoshop Extended versions CS3 (version 10) - CS6 (version 13). To
sign up, go to:

www.Imagingandanalysis.com/web082012ms.html.

You will learn to post-process movie files and things like putting
movies side-by-side, adding arrows and other symbols, adding captions,
and so on. Here is a list of what will be shown:

- Open Image stacks as image sequences; or as a z- or time-series saved
in the uncompressed AVI format
- Maintain a log of what you did to movie files
- Open DICOM series as movies
- Add text, arrows, captions, etc. to movies
- Filter movies all at once: De-Interlace, Sharpen, etc.
- Place movies side-by-side; overlay movies to merge
- Adjust tonal range for optimizing to output devices
- Export completed movies for frame-by-frame "scientific" replay

NOTE: THERE IS STILL ROOM AVAILABLE FOR THE 3RD ANNUAL SCIENTIFIC
IMAGING & QUANTITATION WORKSHOP. See imagingandanalysis.com.

--
Jerry (Gerald) Sedgewick
Scientific Imaging & Image Analysis Consultant
Technical Writer / Regulatory Consultant
965 Cromwell Avenue
Saint Paul, MN 55114
651-788-2261
http://www.imagingandanalysis.com
http://www.quickphotoshop.com
Author: �Scientific Imaging with Photoshop: Methods, Measurement and Output�

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LRW certainly doesn't autofluoresce and I doubt Unicryl does. I assume you are sticking to formaldehyde or formalin - glutaraldehyde will cause autofluorescence. For LM, I prefer butyl methyl methacrylate (BMMA) since you can extract the resin with acetone and it is great for immunocytochemistry.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

} realname - Matt
} Email - m1graus-at-gmail.com
} ORGANIZATION - University of New Mexico EDUCATION - Graduate College
} SUBJECT_OF_QUESTION - auto fluorescence QUESTION - I'm trying to embed
} yeast cells into both unicryl and LR-white.� My question is do either
} media auto fluoresce?

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I'm hoping that someone out there may have a good set of schematics for a
Cambridge S-90 and particularly the framestore option. I'd be willing to
pay any reasonable cost for digitizing/copying or make arrangements for it.
This is for a customer who can't afford a newer instrument but needs to keep
this old one running. It was bought used and didn't include a schematic set
and apparently the manufacturer can no longer provide a useable set.

Allen R. Sampson
Owner
Advanced Research Systems
317 North 4th. Street
Saint Charles, Illinois �60174
ars-at-advressys.com �ph (800) 506-9770

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Dear all,
My TEM H-600 has image distoration possiblely due to contamination in the column. Please see picutre below
http://farm9.staticflickr.com/8299/7771051022_74b6521371_m.jpg

Can anyone help me to point out which part is contaminated, so I can take that part out and clean it.

Thank you very much!

Gary Zhang

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Hi

I may be able to help you but your link to a picture ended with "This photo
is currently unavailable?"

The most common places for "dirt" problems in a TEM are the condenser
system, the specimen rod and the area just below the specimen position. A
problem in the area just below the specimen will give you chronic image
astigmatism and problems any further down the column will give an image
distortion. My guess is that if this is a flicker that varies with
adjustment of condenser 2 (brightness) you need to clean the area between
this lens and the bottom of the illumination alignment coils. But to
isolate the problem further try the following.

1. Lower the accelerating voltage which should increase the problem,
raising the voltage the problem should be reduced. If not the latter the
problem is a dirty gun chamber; physical dirt or poor vacuum.
2. Change the spot size - larger (bigger problem) or smaller (lesser
problem) this will confirm that the problem is below the first condenser
lens pole piece.
3. Similarly change the condenser aperture - larger (bigger problem) or
smaller (lesser problem) this will confirm that the problem is below the
condenser aperture.

Hope this helps if not come back to me with the results of the above tests.

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
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To: protrain-at-emcourses.com

Dear all,
My TEM H-600 has image distoration possiblely due to
contamination in the column. Please see picutre below
http://farm9.staticflickr.com/8299/7771051022_74b6521371_m.jpg

Can anyone help me to point out which part is contaminated, so I
can take that part out and clean it.

Thank you very much!

Gary Zhang

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I have traditionally used Reynold's lead citrate made from lead nitrate but Venable & Coggeshall's formula that uses solid lead citrate as a starting material always seemed to me a better idea. I decided to give the V&C protocol a shot but my first two attempts have failed. I used freshly prepared NaOH stocks made from boiled deionized water and carbonate-free concentrated 50% NaOH solutions. I used "carbonate free lead citrate" solid from a major EM supply house (albeit a bottle that was several years old). V&C suggest 0.01 to 0.04 g lead citrate in 10 mls of 0.01 M NaOH and I used 0.02 mg per 10 mls. In both my attempts, most but not all the solid goes into solution. This is naturally unnerving. I am about to revert to Reynold's formulation unless someone has thoughts on what is going wrong. I am aware of other formulations like Sato, etc but really want to restrict my efforts to V&C or Reynold's. Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Hi Thomas,
I have tried both methods. The solid lead citrate has never worked for us. Not sure why that is since like you we have made it with exact detail. No matter what we did, boil water, make fresh NaOH, sit inside a vacuum (HaHa) we always got a tremendous amount of precipitate. So we just stick to the plain ole Reynold's recipe and fortunately that works 95% of the time.

Lita Duraine
EM Technologist
Bellen Lab
HHMI- Molecular Genetics
Duncan Neurological Research Institute
1250 Moursund St.
Houston, TX 77030
Rm: N1165.17
MS: N1125.50
832-824-8772 TEM Room
979-549-6526 Cell
http://flypush.imgen.bcm.tmc.edu/lab/people/lita.php

-----Original Message-----
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Sent: Monday, August 13, 2012 1:51 PM
To: Duraine, Lita

I have traditionally used Reynold's lead citrate made from lead nitrate but Venable & Coggeshall's formula that uses solid lead citrate as a starting material always seemed to me a better idea. I decided to give the V&C protocol a shot but my first two attempts have failed. I used freshly prepared NaOH stocks made from boiled deionized water and carbonate-free concentrated 50% NaOH solutions. I used "carbonate free lead citrate" solid from a major EM supply house (albeit a bottle that was several years old). V&C suggest 0.01 to 0.04 g lead citrate in 10 mls of 0.01 M NaOH and I used 0.02 mg per 10 mls. In both my attempts, most but not all the solid goes into solution. This is naturally unnerving. I am about to revert to Reynold's formulation unless someone has thoughts on what is going wrong. I am aware of other formulations like Sato, etc but really want to restrict my efforts to V&C or Reynold's. Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

==============================Original Headers==============================
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I decided the uncertainty wasn't worth it and made a batch of Reynold's lead citrate. It worked perfectly with the NaOH stocks I had. I appreciate all the comments I got both on list and off list. It appears I am not alone in having problems making the V&C method work. Now I need everyone to think good thoughts as I go over to the TEM and look at my grids later this week; I need all the positive vibes you can spare. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Email: dl52-at-duke.edu
Name: D. N. Leonard

Organization: Evans Analytical Group

Title-Subject: [Filtered] AReMS Fall 2012 Meeting Announcement

Message: Greetings Microscopy Colleagues,

The Appalachian Regional Microscopy Society (AReMS) invites you to attend the upcoming Fall 2012
meeting being held Oct. 11-12 in Knoxville, TN! Meeting registration ($70 regular, $35 student) will
include a FULL DAY of hands on tutorials, a meeting banquet ticket, presentations on
state-of-the-art microscopy by experts in the field and a tour of Oak Ridge National Laboratory's
SNS, CNMS and ShaRE facilities. The conference hotel is just a 5 minute walk to the Univ. of Tenn.
electron microscopy center and the meeting banquet is being held at Calhoun's on the River. Several
vendors will be in attendance and available to provide information about new products or answer
questions. More information about the meeting can be found below.

- Visit the AReMS website www.arems.org to register (Paypal accepted), submit an abstract and find
information about the conference hotel

- Deadline for student awards, poster abstract submission and ORNL tour is Sept. 7, 2012.

AReMS is very supportive of student participation and there are always expert microscopists in
attendance who are fun to socialize with and can answer specific microscopy related questions. We
look forward to seeing you in Knoxville!

Cordially,
Donovan N. Leonard
President and Program Chair
Appalachian Regional Microscopy Society
www.arems.org
info-at-arems.org
---------------------------------------------------------------------------------------------------------------------------------------------------------

Presentations will be given by:

Dr. David Joy (UTK/ORNL)
Dr. Stephen Pennycook (ORNL)
Dr. Dave Piston (MSA past-president MSA Tour Speaker, Vanderbilt)
Dr. John Henry Scott (MAS Tour Speaker, NIST)
Dr. James LeBeau (NCSU)
Dr. Jeffrey Caplan (Del. Biotech. Inst.)

Tutorials being offered include:

- Correlative microscopy: correlating back scattered TEM-like SEM images and super resolution microscopy

- Zeiss Libra200 will be used in STEM mode to show how a monochromator can be applied to improve
EELS energy resolution for a plasmonics application.

- Zeiss Libra200 STEM/EELS core-loss acquisition and data processing will be demonstrated.

- Zeiss applications engineer will be present to demo the Auriga SEM/FIB capabilties as applied to
biological materials imaging and FIB slicing.

- Zeiss applications engineer will be present to demo the Auriga SEM/FIB capabilties and review the
step by step process of preparing a TEM specimen using liftout and FIB thinning.

- Cryo-ultramicrotomy sample preparation of polymers, for cryo-stage TEM analysis, will be
demonstrated by Leica.

- Cryo-ultramicrotomy sample preparation of a biological material, for cryo-stage TEM analysis, will
be demonstrated by Leica.

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Email: kwilliam-at-mun.ca
Name: Kate Williams

Organization: Memorial University St.John's NL

Title-Subject: [Filtered] servicing ultramicrotomes

Message: We have two Leica ultra microtomes an ultracut E and ultracut S that we would like to have
serviced.
I have asked Leica but they are no longer servicing these models.
Can anyone recommend someone who can service these models? (We are located in Newfoundland eastern
Canada)
Thank you
Kate

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Email: stacie-at-ems-secure.com
Name: Stacie Kirsch

Organization: Electron Microscopy Sciences

Title-Subject: [Filtered] ImmunoGold Fall Workshop

Message: Aurion and Electron Microscopy Sciences are pleased to announce the Fall Workshop, Aurion
ImmunoGold Silver Staining, to be held at the University of Minnesota, Minneapolis Characterization
Facility October 8-10, 2012. For more information, visit
http://www.emsdiasum.com/microscopy/products/immunogold/workshop.aspx or contact me directly.

Thank you,
Stacie Kirsch
Electron Microscopy Sciences
1560 Industry Road
Hatfield, PA 19440
Tel:215-412-8400
Fax:215-412-8450
E-Mail:sgkcck-at-aol.com or stacie-at-ems-secure.com
Web:www.emsdiasum.com

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Email: m.aqel-at-mail.utoronto.ca
Name: M.Aqel

Organization: University of Toronto

Title-Subject: [Filtered] OXFORD INCA x-sight

Message: Hi

anyone running ISIS 1.04 software with Oxford Inca x-sight under windows 3.11 for workgroup?
I have few questions to ask

Thank you
M.Aqel

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Email: wadowska-at-upei.ca
Name: Dorota Wadowska

Organization: Atlantic Veterinay College/UPEI

Title-Subject: [Filtered] EM textbook

Message: Last year I heard that the third edition of Bozzola and Russell Electron Microscopy
principles and techniques for biologists was going to be published. Timing was fall 2011. I did a
search on Internet and I could not find any 3rd edition of this book. Do you know if it was
published and if yes who was the publisher? Could you recommend any newer textbooks?
Thank you
Dorota

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Email: derek.hammill-at-carestream.com
Name: Derek

Organization: Carestream Health

Title-Subject: [Filtered] SEM Resolution

Message: I am slowly loosing resolution in my SEM images over the course of a few years. We have on
occasion placed oils/greases in there for analysis as well as other materials which may outgas. Is
this likely my problem? What can I do now?

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Email: kaszas.1-at-osu.edu
Name: Andrea Kaszas

Organization: Ohio State University/OARDC

Title-Subject: [Filtered] Sputter coater

Message: We have been using the Hummer VII instrument for coating our SEM samples for over 20 years.
This instrument recently broke and we can not repair it. We like Hummer VII spatter coater because
of its ease of use and fast processing time. As they don't make this model any more, would anyone
know which instrument would the most similar to this one? Or, can you suggest an instrument for this
purpose?

Thank you.

Andrea Kaszas

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X-from: Gregory.Hendricks-at-umassmed.edu ()

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Email: Gregory.Hendricks-at-umassmed.edu
Name: Greg Hendricks

Organization: UMass Medical School

Title-Subject: [Filtered] Research Associate Position Opening

Message: Description:

The Core Electron Microscopy Facility at the University of Massachusetts Medical School seeks a
highly motivated individual with significant experience in biological electron microscopy.
Responsibilities will include operation of multiple Transmission Electron Microscopes and a new
state-of-the art Field Emission Scanning Electron Microscope (FESEM), together with user training
and support. As part of a growing, world-class electron microscopy user facility, the successful
candidate must be able to communicate effectively with faculty and students, and have demonstrated
ability in specimen preparation, including fixation, embedding and sectioning of routine specimens
as well as samples prepared for immune-EM. He/she will help in training users in microscopy
techniques and microscope operation.

Qualifications:

Candidates will have a Bachelors or Master's degree, or the equivalent education, and a minimum of
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individual should have significant practical experience in EM in general and a sound understanding
of the associated analytic techniques such as EDS.
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The University of Massachusetts Medical School is an equal opportunity, affirmative action educator
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Contact: Gregory Hendricks, PhD
Manager, Core Electron Microscopy Facility
University of Massachusetts Medical School
Gregory.Hendricks-at-umassmed.edu

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Hi Derek,
There is a device specifically designed to clean residual hydrocarbons from
Electron Microscopes called a "downstream plasma cleaner". This puts Oxygen
radicals into the microscope, they react with the hydrocarbon contamination,
volatilize it, and the volatiles are pumped away, restoring instrument
performance. I don't know where you're based but you can find XEI Scientific
Inc for more details.

est Regards,

Ian Holton
Acutance Scientific

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Email: derek.hammill-at-carestream.com
Name: Derek

Organization: Carestream Health

Title-Subject: [Filtered] SEM Resolution

Message: I am slowly loosing resolution in my SEM images over the course of
a few years. We have on occasion placed oils/greases in there for analysis
as well as other materials which may outgas. Is this likely my problem?
What can I do now?

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Good morning/good afternoon,
dear Prof. Phillips, dear all,

Apologize if I am chiming in lately with this...
besides I would really be glad to receive also the offline responses,
as (only) a {satisfied} long-standing user of Venable & Coggeshall's Lead Citrate solution (no commercial or financial interest) I would like to comment on the making and use of the solution.

{ {I decided to give the V&C protocol a shot but my first two attempts have failed.} }

{ {I used freshly prepared NaOH stocks made from boiled deionized water and carbonate-free concentrated 50% NaOH solutions.} }

The original recipe says:
^^^^^^To make 10 ml of the stain, simply weigh out 0.01 to 0.04 grams of lead citrate, add it to 10 ml of singly distilled water in a screw-topped centrifuge tube, then drop in 0.1 ml of 10 N sodium hydroxide. Close the tube tightly and shake it vigorously until all the lead citrate is dissolved. The total operation takes less than 5 minutes. - ....... -
To prevent the formation of lead carbonate as a precipitate, avoid contaminating the staining solution with carbonate ions. Use only concentrated sodium hydroxide solutions or sodium hydroxide pellets in making the stain since these forms contain little sodium carbonate. Also, keep the tube sealed from atmospheric carbon dioxide.
A lengthy exposure to the atmosphere must occur before precipitates render the stain useless.
Stain sections by placing them, while on supporting grids, either in or upon small quantities of the staining solution dropped on clean wax or glass. Staining time required for Epon or Araldite sections varies from 10 seconds to 5 minutes, depending upon the type of tissue and its previous treatment. - ...... -^^^^^^

Comment: IMHO it is the altered mixing procedure that makes you "itching" / troubles....

{ {I used "carbonate free lead citrate" solid from a major EM supply house (albeit a bottle that was several years old).} }
Comment see below (next statement)

{ {V&C suggest 0.01 to 0.04 g lead citrate in 10 mls of 0.01 M NaOH and I used 0.02 mg per 10 mls.} }
Comment: according to the original recipe, 0.01-0.04 lead citrate (from K&K Labs, what the properties of this stuff really were in 1965, we unfortunately don't know today) is added to single (or better: double) distilled water...see above.
I always (since 1981 the same stuff) used lead(II)citrate-3-hydrate (C6 H5 O7)2 Pb3.3H2O (e.g. MERCK #12438,50 or 100g). 0.01 g as well as 0.04 g is not quite easy to weigh (if you don't have a microgram-balance at hand).When I did this several times -seeing the "real amount"- I decided just to get a pinch of Pb-citrate-"granules" out of the plastic container. The assumptive "real" amount of Pb-citrate powder will be between 0.010-0.025 g, which is sufficient. IMO, it is necessary to dissolve (at least a little bit!) the Pb-powder first in "plain" hot A. bidest, NOT in 0.01 M NaOH.

My procedure is as follows (all components ready on the table/preparation site):
i) boil approx. 100 ml A. bidest. (if NOT freshly/hot from the distilling apparatus) for at least 2 min to get rid of eventually present CO2 in the water.
ii) with most of this volume (70-80 ml) boil a PRE-CALIBRATED (make a 10 ml-mark for hot water before) clean snap cap glass vial and its snap cap lid too (1-2 min)
iii) Fill the {hot} glass vial up to the 10 ml-mark with the remaining boiling/boiled/hot A. bidest.
iv) get a pinch / mote of crystalline Pb-citrate out of the container (be careful in not wetting the mote of powder by water vapor coming out of the vial) by a clean spatula and place it into the HOT A. bidest. in the vial.
v) add 100 �l of 10 N NaOH (use Micropipette) to the solution (I actually use a solution which has been made in stock in 2002, but stored in a plastic bottle with plastic stopper on the Lab-shelf), then immediately
vi) close the glass vial with the plastic snap cap lid and
vii) shake vigorously (use safety gloves protecting your fingers against the heat!) at least for 2-4 min.
viii) have a look for white powder remnants, if yes: don't open the plastic lid totally from the vial but instead lift off a little bit and add additional 100 �l of 10 N NaOH (40g NaOH/100ml A. bidest.)....the pH of nearly 12 cannot be increased with the additional amount of NaOH but perhaps solves the remaining white "precipitate" after further vigorous shaking of the vial.
IF NO white precipitate was found: you can cool the vial under tap water to RT.

Tip: How to use & When using:
i) always keep the solution in the dark (Al-foil wrapping) for storage,
ii) just lift the plastic snap cap only a little bit and suck some 100 �l or as needed for four to five drops into a micropipette which has been swilled at least once with NaOH solution before.
iii) position drops for Pb-staining as usual in a wax-layered Petri dish (+ NaOH-pellets round about).

Staining time: IMHO will depend on resin used and the previous uranyl-acetate staining, as well as the age of the Pb-citrate solution made and the "quality" of the glass of the vial used.
We/I have found that with a freshly produced Pb-stain, staining time should not exceed 30 seconds, if the solution stood for several days staining time can be increasingly longer (say 1d - 5d = 1-1.5 min, 5-15 days: up to 3 min, depending also how often the vial was opened) which perhaps is correlated with a modest reaction between the highly alkaline Pb-citrate solution and the glass as well as CO2 uptake despite the solution nevertheless is quite stable for a longer time (up to 4 weeks).
(Pre-Staining of our ultrathins: 5-8 min 0.05% hydrous, filtered (Low Mol. Weight) Tannic acid; followed by routinely 15 min 1% methanolic uranyl-acetate additionally acidified by some drop of conc. acetic acid, light-shielded/in the dark).
In most (98%) of our ultrathins the Pb-contrast is pretty intensive (and - by the way - due to TA-prestaining - also collagen and glycogen will exhibit "specific" heavy electron density).

{ {In both my attempts, most but not all the solid goes into solution.
This is naturally unnerving. I am about to revert to Reynold's formulation unless someone has thoughts on what is going wrong.} }

This is deploring but "Rome wasn't built in a day either" (see above).
In the 1980ies when I started in the EM Lab here I have always done both methods in parallel just to be on the safe side....After the third or fourth time with success I switched exclusively to Venable & Coggeshall...

VENABLE J.H, Coggeshall R. (1965): A Simplified Lead Citrate Stain for Use in Electron Microscopy; J. Cell Biol.(N.Y.) 25, 407-408
[The SCI� indicates that this paper was cited 2,120 times in the period 1961-1975. The .pdf available on request.]
see also:
R. L. RICHARDSON, D. M. HINTON and D. R. CAMPION (1983)
An Improved Method for Storing and Using Stains in Electron Microscopy
J. Electron Microsc., Vol. 32, No. 3, 216-218, ( Storage up to 6 months, a .pdf available on request)

Best wishes and compliments,
Wolfgang MUSS (Member of MSA)
SALZBURG, AUSTRIA

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Gesendet: Montag, 13. August 2012 23:23
An: Mu� Wolfgang
Betreff: [Microscopy] More on lead citrate woes
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I decided the uncertainty wasn't worth it and made a batch of Reynold's
lead citrate. It worked perfectly with the NaOH stocks I had.

I appreciate all the comments I got both on list and off list.
It appears I am not alone in having problems making the V&C method
work.
Now I need everyone to think good thoughts as I go over to the TEM and
look at my grids later this week; I need all the positive vibes you can
spare.
Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
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Lita Duraine
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MS: N1125.50
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http://flypush.imgen.bcm.tmc.edu/lab/people/lita.php

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I have traditionally used Reynold's lead citrate made from lead nitrate but Venable & Coggeshall's formula that uses solid lead citrate as a starting material always seemed to me a better idea.

I decided to give the V&C protocol a shot but my first two attempts have failed.

I used freshly prepared NaOH stocks made from boiled deionized water and carbonate-free concentrated 50% NaOH solutions.

I used "carbonate free lead citrate" solid from a major EM supply house (albeit a bottle that was several years old).

V&C suggest 0.01 to 0.04 g lead citrate in 10 mls of 0.01 M NaOH and I used 0.02 mg per 10 mls.

In both my attempts, most but not all the solid goes into solution.
This is naturally unnerving. I am about to revert to Reynold's formulation unless someone has thoughts on what is going wrong.

I am aware of other formulations like Sato, etc but really want to restrict my efforts to V&C or Reynold's. Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Hello Andrea,
I feel your pain, I grew up (professionally speaking) with that sputter coater. I talked my current company into buying a Cressington 108. I found it easy to operate and works fast. I'm using gold but hope to move to Pd/Pt someday.

Just make sure you buy all the "plumbing" you need. When I purchased mine, I had hopped it would male barb-vacuum hose-male barb. I was wrong.

Stay safe..............
Frank

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Email: kaszas.1-at-osu.edu
Name: Andrea Kaszas

Organization: Ohio State University/OARDC

Title-Subject: [Filtered] Sputter coater

Message: We have been using the Hummer VII instrument for coating our SEM samples for over 20 years.
This instrument recently broke and we can not repair it. We like Hummer VII spatter coater because
of its ease of use and fast processing time. As they don't make this model any more, would anyone
know which instrument would the most similar to this one? Or, can you suggest an instrument for this
purpose?

Thank you.

Andrea Kaszas

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Dorota
I am also waiting for the release of the Bozzola EM textbook 3rd
edition as I use it as a student textbook in my graduate TEM course.
It's more than a year overdue since the promised release date and I'm
not sure of the hold up. Maybe the author has more info?

Tony Greco

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} realname - alaa afeef
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} ORGANIZATION - Glasgow University
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} SUBJECT_OF_QUESTION - Learning about 3D Image Reconstruction in
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} QUESTION - Hello,
}
} I am trying to learn about the Reconstruction Algorithms in Electron
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Email: patrick.trimby-at-sydney.edu.au
Name: Pat Trimby

Organization: The University of Sydney

Title-Subject: [Filtered] FIB-SEM GIS Precursors

Message: Hi Everyone,

We have 2 FIB-SEMs in our lab (one FEI, one Zeiss), and I have a health and safety question relating
to the precursors used with the Gas Injection Systems on these instruments. We are intending to use
XeF2 and WF6, both of which are classed as highly toxic.
At present, the pump system for the FIB-SEMs vent directly into the lab, with no external exhaust
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(and subsequently being pumped into the lab) is extremely small, we are concerned about the use of
these materials. We know of one lab where they were forced by health and safety regulations to
install an exhaust system that vented to the roof of the building, and I wondered if anyone else has
experience in managing the risk with these (and other) toxic gases? As far as I can tell, the
manufacturers do not give any health and safety recommendations for using such precursors (possibly
so as not to dampen sales?!).

Any comments on this issue are welcome.

Regards,

Pat Trimby

The Australian Centre for Microscopy & Microanalysis
The University of Sydney

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Derek,
That is likely the problem. A thorough cleaning of the column from top to
bottom should make a significant difference. The downstream plasma cleaner
might also help.

One other thing to look at is the secondary detector. Are you changing the
scintillator regularly? Does it have a brownish cast to it? Anything that
lessens the contribution of the lowest energy secondary electrons (whether a
dead layer from bombardment or a layer of contamination) first and foremost
cuts into resolution, often without reducing overall signal by much. The
finest details generally come from the lowest energy secondaries.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

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Name: Derek

Organization: Carestream Health

Title-Subject: [Filtered] SEM Resolution

Message: I am slowly loosing resolution in my SEM images over the course of
a few years. We have on
occasion placed oils/greases in there for analysis as well as other
materials which may outgas. Is
this likely my problem? What can I do now?

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Hi Derek

You have had some good replies but may I add my ideas?

In a similar circumstance I have spent a happy hour or two with a hot air
blower heating up the pumping manifold and the specimen chamber. Sounds a
bit of a joke but I can assure you it vastly improved the situation. In
truth the pumping line to the rotary pump should be changed if a really good
job is the object. I automatically assume that in your situation the RP
fluid is changed at least every 6 months?

Enjoy

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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Name: Derek

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Title-Subject: [Filtered] SEM Resolution

Message: I am slowly loosing resolution in my SEM images over the course of
a few years. We have on occasion placed oils/greases in there for analysis
as well as other materials which may outgas. Is this likely my problem?
What can I do now?

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Hi Pat,

This is rather anecdotal as I do not have any supporting documentation,
but I am aware that in the old days of introducing Circuit Edit FIBs
with heavy usage of XeF2 and W(CO)6 to semiconductor FABs one of the
major semiconductor companies did a thorough testing of the FIB exhaust.
As far as I can recall, testing was done of the system with dual-stage
oil-sealed roughing pumps and there were no any traces of precursors or
byproducts detected in the exhaust. The hand-waiving explanation was
that trace amounts of precursors are easily absorbed/dissolved in oil
within roughing pump. I do not know if mineral or synthetic oil was used
during testing, I do not know if such testing was ever done for WF6, and
I am not aware if anyone did such investigation on the system with dry
roughing pumps.

If you have or can rent RGA unit, then you should be able to test
exhaust gas of your FIB system and determine if any precursors or
byproducts are getting out of it.

That being said - I personally do not believe that it may be healthy to
breath for prolonged periods of time anything that we were not
"designed" to breath, even if that is only trace amounts of supposedly
inert vacuum oils. So why not play it safe and exhaust that pump to
outside, just in case? If for any reason external exhaust is unfeasible,
then it is easy and less expensive to install oil mist filer followed by
activated charcoal filter (don't forget to replace cartridge regularly)
on the exhaust of your roughing pump - that would take care of pump oils
and anything else which may or may not come out.

Cheers :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 8/14/2012 9:22 AM, microscopylistserver-noreply-at-microscopy.com wrote:
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}
} Title-Subject: [Filtered] FIB-SEM GIS Precursors
}
} Message: Hi Everyone,
}
} We have 2 FIB-SEMs in our lab (one FEI, one Zeiss), and I have a health and safety question relating
} to the precursors used with the Gas Injection Systems on these instruments. We are intending to use
} XeF2 and WF6, both of which are classed as highly toxic.
} At present, the pump system for the FIB-SEMs vent directly into the lab, with no external exhaust
} system. Although we understand that the likelihood of a whole precursor outgassing in a short time
} (and subsequently being pumped into the lab) is extremely small, we are concerned about the use of
} these materials. We know of one lab where they were forced by health and safety regulations to
} install an exhaust system that vented to the roof of the building, and I wondered if anyone else has
} experience in managing the risk with these (and other) toxic gases? As far as I can tell, the
} manufacturers do not give any health and safety recommendations for using such precursors (possibly
} so as not to dampen sales?!).
}
} Any comments on this issue are welcome.
}
} Regards,
}
} Pat Trimby
}
} The Australian Centre for Microscopy & Microanalysis
} The University of Sydney
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The Institute for Imaging and Analytical Technologies at Mississippi
State University has an opening for a Postdoctoral Research Associate
with expertise in Transmission Electron Microscopy techniques as applied
to the area of physical metallurgy.

https://www.jobs.msstate.edu PARF# 6559

See below for details:

Position Description
Postdoctoral Research Associate
Institute for Imaging & Analytical Technologies
Mississippi State University

In order to properly serve the needs of the growing materials science
and engineering research community at Mississippi State University
(MSU), the Institute for Imaging & Analytical Technologies (I2AT) has an
opening for a Postdoctoral Research Associate with expertise in
Transmission Electron Microscopy techniques as applied to the area of
physical metallurgy. This Postdoctoral Research Associate will report to
the Director of the I2AT, a university-wide core facility and research
institute meeting all three missions of the university, research,
teaching, and service.

Characteristic Duties and Responsibilities
The applicant will be responsible for operation, training,
consultation, scheduling, and routine maintenance of new
state-of-the-art multi-user analytical transmission electron microscope
within MSU*s I2AT. The applicant will primarily support TEM
applications, but will also have the opportunity to learn/operate other
imaging and characterization technologies (e.g. scanning electron
microscopies, X-ray diffraction, scanning probe microscopy) available at
the I2AT. In terms of professional development, the applicant will be
encouraged to develop an independent research program, contribute to
interdisciplinary research efforts, and direct graduate and
undergraduate student research assistants. The applicant will also be
expected to participate in the preparation of peer-reviewed journal
articles, research proposals, and reports.

Minimum Acceptable Qualifications
The successful applicant must have a Ph.D. (or equivalent degree) in
any engineering or physical science field and a strong background in
materials characterization, imaging and diffraction techniques, EM
theory and operation. Knowledge of other general imaging/analysis
approaches is desired. The applicant must have a demonstrated ability to
work independently, to work in a collaborative environment with a
diverse group of colleagues, and to communicate effectively in written
and oral formats.

Additional Information
This is a one-three year, 75% time appointment, leaving an opportunity
for the post-doctoral associate to obtain a 25% appointment from
teaching assignments or extramural research funding. Review of
applications will begin immediately and will continue until the position
is filled.
Applicants should complete an online application at
https://www.jobs.msstate.edu/ and upload a cover letter and current
vitae. Applicants should also submit three letters of recommendation
(mailed by the individual recommending the applicant under separate
cover) and an official complete graduate school transcript. Letters of
recommendation and transcripts should be mailed to Shauncey Hill,
Business Manager, Institute for Imaging & Analytical Technologies,
Mississippi State University, Box 6020, Mississippi State, MS 39762
(shill-at-i2at.msstate.edu, 662-325-8739).

==============================Original Headers==============================
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Hi Alaa,

Check out the Boluder Lab for info.

The Boulder laboratory for 3-D electron microscopy of cells has functioned as a NCRR funded research resource since 1970. The lab has been successful in developing methods for imaging diverse cells and tissues in three dimensions (3-D) at a resolution of ~ 5 nm. The numerous features of cell structure that become visible at this level are highly informative about the mechanisms that underlie cellular processes.

http://bio3d.colorado.edu/

Roseann

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

On Aug 14, 2012, at 8:21 AM, oshel1pe-at-cmich.edu wrote:

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} } realname - alaa afeef
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} } ORGANIZATION - Glasgow University
} } EDUCATION - Graduate College
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} } SUBJECT_OF_QUESTION - Learning about 3D Image Reconstruction in
} } Electron tomography
} } QUESTION - Hello,
} }
} } I am trying to learn about the Reconstruction Algorithms in Electron
} } Tomography, Unfortunately I dont know from where I should start as
} } most of the books out there is about CT reconstruction.
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Email: gul417-at-mail.usask.ca
Name: Guosheng Liu

Organization: U of Saskatchewan

Title-Subject: [Filtered] Re: servicing ultramicrotomes

Message: Hi Kate,

Chris did an excellent job for our ultracut E last year. He has moved in Calgary I guess. His
contact info is:

Chris Cathcart
Prismatic Solutions
647.975.5210

Good luck

Guosheng Liu

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Alaa:

Regarding Electron Tomography, I too checked the ever useful Matlab Users
forum. And, while I did not find code for electron tomography there is code
for Xray reconstruction, acoustic reconstruction, .... The mathematics of
reconstruction is not that different once images are generated, in my
opinion. I would definitely start with Matlab file exchange, read the code
look at common elements between the Xray and Sonic algorithms then change
the code slightly for the Physics of electrons. Good luck!

http://www.mathworks.com/matlabcentral/fileexchange/?search_submit=fileexcha
nge&query=Tomography&term=Tomography

Pete Eschbach
Former TEM guru at Fortune 30 company :o)

-----Original Message-----
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***********
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} realname - alaa afeef
} Email - alaa.afeef-at-gmail.com
} ORGANIZATION - Glasgow University
} EDUCATION - Graduate College
} LOCATION - Glasgow
} SUBJECT_OF_QUESTION - Learning about 3D Image Reconstruction in
} Electron tomography
} QUESTION - Hello,
}
} I am trying to learn about the Reconstruction Algorithms in Electron
} Tomography, Unfortunately I dont know from where I should start as
} most of the books out there is about CT reconstruction.
}
} If I may to, ask for your help in directing me about this issue,
} recommending some Introductory readings and MATLAB simulations if
} any.
}
} Thanks you very much in advance for your help

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Hi

The picture that you sent is typical of dirt in the imaging system
exhibiting charge! It is very unusual for this to suddenly happen and I
believe that you may have a lens that is not working. Why I say this is
that the image you have is typical of viewing an area that you do not
normally visualise when the lenses are working correctly.

I suggest you take note of the magnification setting when the problem is
first visualised and turn the magnification down one step. If the problem
goes away my theory is correct; you have lost a lens. If you area able to
see the lens currents as you make these changes you may be able to deduce
which lens is at fault. I would expect the problem to be a power transistor
in that lens circuit as they are the most likely component to fail.

Please come back to me if you need more.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: Gary Zhang [mailto:zhanggary717-at-yahoo.com]
Sent: 15 August 2012 08:22
To: protrain-at-emcourses.com

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Email: grzegorzpolak-at-mac.com
Name: Grzegorz (Greg) Polak

Organization: Hofstra University

Title-Subject: [Filtered] Specific TEM Stains

Message: I am currently involved in the research and Hofstra University graduate program. I'm
working on my masters degree and part of my research is use of TEM microscope to look for specific
glycolytic enzymes that are found/missing on the myofibers in a specific muscles Mio (ChREBP)
knockdown in Drosophila. I know that there are specific immunostain's for most of the glycolytic
enzymes but because I'm looking into muscle specifically I would like to see if there is any
distribution changes and placement of these enzymes in the muscle. Since I'm taking a picture with a
TEM to see if there's a difference between a Wild type and Mio knockdown in the muscle fiber
morphology, I would also at the same time like to see if there is changes in a glycolytic enzyme
placement.

The question is: are there specific heavy stains that will attach directly to the following enzymes
thus making them detectable and visible to see if there are changes between Wild type and Mio
knockdown flies.
Enzymes are: GPDH, GAPDH,TPI, PGK, Aldolase.

Also can I target Z discs and M-lines directly and if yes with what stains.

Thank you for your time and patience, this is the last step in my research and it seems to be the
hardest so far. I hope I made my project somewhat clear and understandable.

Thanks Greg

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Dear Collective,

Does anyone out there have a pet method for collection and processing of platelets for SEM? I have viewed some online articles and have a pretty good idea about the general way to go, but it's always nice to check with the pros!

Thanks in advance.

Cheers,
Randy

Randy Tindall
Senior Research Specialist
Electron Microscopy Core Facility
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65201
(573) 884-5383

We Do Small Well!

www.emc.missouri.edu (dayjob)
http://nadiasyard.com (blog)
http://www.facebook.com/RandyTindallGalleries (photography)

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Greetings;
I am trying to use Perl's iron stain to look at iron deposits in grains. I did a trial run in February, and got great staining, but have been unable to reproduce it since. Does anyone have experience with using this stain on GMA sections? Any advice would be most welcome.

Thanks in advance

Dr. S. Shea Miller
ECORC | CRECO
Agriculture and Agri-Food Canada | Agriculture et Agroalimentaire Canada
960 Carling Avenue | 960, avenue Carling
Ottawa, ON K1A 0C6
E-mail Address / Adresse courriel shea.miller-at-agr.gc.ca
Telephone | T�l�phone 613-759-1760
Facsimile | T�l�copieur 613-759-1701
Teletypewriter | T�l�imprimeur 613-773-2600
Government of Canada | Gouvernement du Canada

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Greetings Fellow Microscopists,

The Hooke College of Applied Sciences, located in Westmont, IL, is offering a transmission electron microscopy short course October 2-4, 2012.� In addition to lectures, this course emphasizes hands-on training using state-of-the-art equipment.� For further TEM training details and registration information, please follow the link below:

http://www.hookecollege.com/courses/course.asp?COURSE_ID=53

Best regards-
__________________________________________________
Chris Gorman
Admissions Specialist
Hooke College of Applied Sciences
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
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Hello EM Community,

We have been asked to look at the ultrastructure of neurons in the C. elegans nervous system. I was wondering if any of you have experience with fixation and epoxy resin embedding of these organisms and are willing to share your expertise.

Thank you so much!
Sandy Hancock

Sandy Hancock
Laboratory for Neurotoxicity Studies
Phase II, Duckpond Dr.
VMRCVM
Virginia Tech
Blacksburg, VA 24061
Phone: 540-231-4817
Email: hancocksk-at-vt.edu

==============================Original Headers==============================
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7, 23 -- Subject: TEM_fixation for C. elegans nervous system
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Dear Sandy,
I would suggest that you consider HPF + FS, finally embedding in resin (Epon,
Lowi).
Many reviews cover this method for C.elegans, e.g.
Three-Dimensional Reconstruction Methods for Caenorhabditis elegans
Ultrastructure
Thomas Müller-Reichert, Joel Mancuso, Ben Lich, and Kent McDonald (2010)
Methods in Cell Biology Vol 96 (EM of model systems), p.332
kind regards,
Reinhard

--
Prof. Dr. Reinhard Rachel
University of Regensburg
Centre for EM / Anatomy
Faculty of Biology & Preclin. Med.
Universitaetsstrasse 31
D-93053 Regensburg - Germany
tel +49 941 943 2837, 1720
fax +49 941 943 2868
mail reinhard.rachel-at-biologie.uni-regensburg.de
office: VKL 3.1.29

next microscopy conferences:
http://www.emc2012.org.uk/
EMC 2012 in Manchester, UK
http://www.mc2013.de/
MC2013 in Regensburg, Germany

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3, 25 -- From Reinhard.Rachel-at-biologie.uni-regensburg.de Wed Aug 15 15:34:08 2012
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3, 25 -- To: "microscopy listserver" {Microscopy-at-microscopy.com} , {hancocksk-at-vt.edu} ,
3, 25 -- {skperkin-at-vt.edu}
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Hello Sandy,

I agree with prof Rachel and would like to add that you might want to try Self-Pressurized Rapid Freezing as an alternative to the established HFS methods. We could achieve the same structural preservation quality using SPRF even with the straight tube method and published this in J. Microsc. 235 (2009), 25-35 (Leunissen & Yi).

Please do not hesitate to contact me or Hong Yi (hyi-at-emory.edu) off list for details.

All the best,

Jan Leunissen
New Zealand

i: www.aurion.nl

On 16/08/2012, at 8:17 AM, skperkin-at-vt.edu wrote:

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} Hello EM Community,
}
} We have been asked to look at the ultrastructure of neurons in the C. elegans nervous system. I was wondering if any of you have experience with fixation and epoxy resin embedding of these organisms and are willing to share your expertise.
}
} Thank you so much!
} Sandy Hancock
}
} Sandy Hancock
} Laboratory for Neurotoxicity Studies
} Phase II, Duckpond Dr.
} VMRCVM
} Virginia Tech
} Blacksburg, VA 24061
} Phone: 540-231-4817
} Email: hancocksk-at-vt.edu
}
}
}
}
} ==============================Original Headers==============================
} 7, 23 -- From skperkin-at-vt.edu Wed Aug 15 15:15:43 2012
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} 7, 23 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
} 7, 23 -- Date: Wed, 15 Aug 2012 16:15:42 -0400
} 7, 23 -- Subject: TEM_fixation for C. elegans nervous system
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--Please respect nature in and around New Zealand's precious waterways and refrain from jet boating--

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Dear Listers

Our lab is in the process of replacing some diamond knives.
I would be grateful if anyone with experience could give me some feedback on your opinion of the DEYEmond brand, specifically the ULTRA 45, from Scimed Scientific Medical Products.
Please direct you replies to my e-mail address.
Thanking you in advance
Alan Hall

Laboratory for Microscopy & Microanalysis
Room1-33, Natural Sciences 2
University of Pretoria
Lynnwood Road
Pretoria 0002
South Africa
Tel: +27 12 4202075
Fax: +27 12 3625150
Mobile: +27 82 3319040

Laboratory for Microscopy & Microanalysis
Private Bag X20
Hatfield 0028
South Africa

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Hello Sandy:

I have worked with C. Elegans a few times in the past. This first time I
tried fixing the buggers they just would not fix seemingly no matter what
concentrations or combinations of fixs I used (Glut, Formaldehyde, OsO4,
acrolien . . . ) then I tried simply popping them in the microwave for 5-10
seconds and bam! they fixed. I did not use a fancy EM Fixation vented
microwave, I just used a little old one that we use for warming buffers and
melting agar. So: Place worms in vials with aldehyde fix in buffer (room
temp), place in microwave at 100% for 5-10 seconds, continue fixation at
room temp for desired time.

Last time I used 2.5 % Glut in Sodium cacodlyate buffer for confocal. Used
the Glut Autofluoresence for imaging the worms. Worked beautifully, I would
use what ever fix you think would be best for EM.

On 15 Aug 2012 at 16:19, skperkin-at-vt.edu wrote:

}
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} Hello EM Community,
}
} We have been asked to look at the ultrastructure of neurons in the C.
} elegans nervous system. I was wondering if any of you have experience
} with fixation and epoxy resin embedding of these organisms and are
} willing to share your expertise.
}
} Thank you so much!
} Sandy Hancock
}
} Sandy Hancock
} Laboratory for Neurotoxicity Studies
} Phase II, Duckpond Dr.
} VMRCVM
} Virginia Tech
} Blacksburg, VA 24061
} Phone: 540-231-4817
} Email: hancocksk-at-vt.edu
}
}
}
}
} ==============================Original
} Headers============================== 7, 23 -- From skperkin-at-vt.edu
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} 16:15:43 7, 23 -- -0400 7, 23 -- From: "Hancock, Sandy"
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} {Microscopy-at-microscopy.com} 7, 23 -- Date: Wed, 15 Aug 2012 16:15:42
} -0400 7, 23 -- Subject: TEM_fixation for C. elegans nervous system 7,
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Richard E. Edelmann, Ph.D., Director
Center for Advanced Microscopy & Imaging
9C Upham Hall
Miami University, Oxford, OH 45056
Ph: 513.529.5712 Fax: 513.529.4243
E-mail: edelmare-at-muohio.edu
http://www.cami.muohio.edu

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I'm looking from a little help on what should be a simple problem. I'm using an older Iridium Ultra (ver1.3 rev D) EDS system. I just trying to dot map a surface of five elements and I have two elements with the same color.

The retched system will not allow me to change the colors. I've tried their help software, different map pallet controls, nothing... I've very frustrated at my inability to make such an elementary and basic change.

To me their system lacks intuitive flow. Spending an hour trying to change a dot map color seems excessive. I've checked with my co-worker and we tried all of his ideas.

Does anyone have any experience or with this freaking, fracking system?

If any one in Iridium Ultra land is out there and wants to send a quarter to call, I can be reached at 330-794-6600 and ask for Frank in microscopy...........

Stay safe..............

Frank
ARDL

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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We have a hot stage for our Zeiss EVO-50 SEM and are interested in setting it up but do not have the accompanying manual. It is an Ernest F. Fullam No 1820 Substage heater. (note: Fullam is now part of MTI). We would truly appreciate it if anyone in possession of a manual for the stage would be willing to email it to me at alawrence-at-i2at.msstate.edu or let me know how we might acquire a copy of the manual.

Thanks!

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I want to thank all the people who contacted me and gave me advice and assistance.
It was a frustrating hour. Just because I can't see the flow or intuitive logic of the Iridium Ultra system doesn't mean you won't either. Several people told me how much they loved their system. Well, the heart wants what the heart wants....

Hey, after more then 25 years of running electron scopes and EDS systems I'm sure this old dog can learn new tricks. I'm starting with "Play dead, Fido."

In case you have the same system here are the steps that finally worked for me.-----

Assuming you have a labeled spectrum
I selected FAST MAP. Then that comes up, go to
PROPERTIES on the main menu bar.
This will open a window with three tabs. You want the one called ELEMENT SELECT.

This produces a window with a smaller inset window with a scroll bar containing colors, element, line ID, range. You need to DOUBLE CLICK THE COLOR associated with the element you want to recolor.

Another window will open and you'll find a box identified as "SPECTRUM WINDOW COLOR"

Wait, wait you're not home yet. Oh, you can change the color there, but I can't figure out that it does. It does change the color next to the element in the previous window, but your dot map remains the same unwanted color. Don't use that button.

No, you want the button above labeled PSEUDO GRAY. Don't worry you're not changing your map to some gray scale. Click that and a color pallet come up and you can (YES Now! Push the button Igor!!!) finally change the color of your dot map for a selected element.

I suspect there maybe other way to get into the barn without going through Sherwood Forest

Again, let me offer my apologies for any aspersions and doubt I may have cast on the Iridium Ultra EDS system. It's a great way to spend 6 hours.

This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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Greetings Fellow Microscopists,

The Hooke College of Applied Sciences, located in Westmont, IL, is offering a scanning electron microscopy short course October 15-19, 2012.� In addition to lectures, this course emphasizes hands-on training using state-of-the-art equipment.�

For further SEM training details and registration information, please follow the link below:

http://www.hookecollege.com/courses/course.asp?COURSE_ID=42

Best regards-
__________________________________________________
Chris Gorman
Admissions Specialist
Hooke College of Applied Sciences
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7412(fax)
www.hookecollege.com

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Hello fellow microscopists (in this case especially the more senior ones!),

I have an old TEM test sample on a grid that I would appreciate help in identifying.

It is a Polaron 0316 PG-IR specimen. Anyone know what it is?

Thanks,
Louie

--

Louis Kerr
lkerr-at-mbl.edu

Research and Education Support Coordinator
Marine Biological Laboratory
7 MBL Street
Woods Hole, MA 02543
508-289-7273
508-540-6902 (FAX)
508-292-0289 (Cell phone)

VISIT OUR WEB SITES:
http://www.mbl.edu/
http://www.courses.mbl.edu/

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Hi

I think you will find it is a Pt-Ir grid?

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: lkerr-at-mbl.edu [mailto:lkerr-at-mbl.edu]
Sent: 17 August 2012 22:22
To: protrain-at-emcourses.com

Hello fellow microscopists (in this case especially the more senior ones!),

I have an old TEM test sample on a grid that I would appreciate help in
identifying.

It is a Polaron 0316 PG-IR specimen. Anyone know what it is?

Thanks,
Louie

--

Louis Kerr
lkerr-at-mbl.edu

Research and Education Support Coordinator Marine Biological Laboratory
7 MBL Street
Woods Hole, MA 02543
508-289-7273
508-540-6902 (FAX)
508-292-0289 (Cell phone)

VISIT OUR WEB SITES:
http://www.mbl.edu/
http://www.courses.mbl.edu/

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Geochronology/Geochemistry/Petrology/Mineral Physics
The Department of Earth Science at the University of California at
Santa Barbara seeks a broadly educated geoscientist who conducts
creative research on the long-term evolution of the solid Earth. A
strong field orientation combined with expertise in analytical tools,
such as electron-probe micro-analysis, electron-backscatter
diffraction or mass spectrometry, is required. The appointee is
expected to develop a vigorous, externally funded research program and
teach a broad spectrum of undergraduate and graduate courses. This
tenure-track appointment will be as an Assistant Professor to begin
July 1, 2013.
A Ph.D. is required at the time of appointment. Review of applications
will begin October 15, 2012. Applicants should request three referees
to send letters of evaluation by October 15. Applicants should submit
a PDF containing a letter of application, curriculum vita, a
description of teaching and research objectives and accomplishments,
and the contact information of the referees who are providing letters.
The application file and letters of reference should be submitted to
betancourt-at-geol.ucsb.edu. Queries about this application can be
directed to Bradley Hacker (hacker-at-geol.ucsb.edu).
The department is especially interested in candidates who can
contribute to the diversity and excellence of the academic community
through research, teaching and service. For more information about the
department, visit our webpage (www.geol.ucsb.edu). UCSB is an Equal
Opportunity/Affirmative Action employer

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Email: ewood1-at-uvm.edu
Name: Erin Wood

Organization: University of Vermont-School of Engineering

Title-Subject: [Filtered] JEOL JSM-6060 SEM jet assembly

Message: Hello everyone,

I hope this inquiry reaches you all in good health and spirits! I am a graduate student in the
mechanical engineering program at the University of Vermont.
I am curious if anyone has any experience with JEOL diffusion pumps, specifically, I believe the 4
inch pump. It seems that our diffusion pump maybe had some contamination in it so we tried to take
it apart to clean, and the aluminium shoulder screws at the bottom got stuck in the little Y shaped
connector in the jet assembly. Long story short, we are at the point where we will now try to drill
out the Y-shaped connector and retap it, and get some new shoulder screws. If this fails we are
looking for a back up plan, so I thought as a last ditch effort we would see if anyone had this
piece from a jet assembly that perhaps is no longer able to be used. This would be easier than
buying the entire jet assembly, as ours is reconditioned and ready to go we think.

We are also curious if the assembly is sitting slightly lower or higher due to the new screws and
threading, will this affect our pumping at all? I'm not sure that contamination was actually the
problem, but it is our best guess that it was. But I'd hate to put the pump back together and have
it not work because it was sitting slightly off from the original position.

Thank you so much for reading this, and for any advice you may offer.

Erin L Wood

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I discovered the other day that several of my
friends who are frequently involved in building
and repairing gadgets in the lab and around home
were not familiar with the PC-7 (black) and
PC-11 (white) epoxy compounds manufactured by the
Protective Coatings Company. These are sold as a
two-component system consisting of a resin paste
and a hardener paste, which are mixed in equal
amounts. They give you about 45 minutes of
working time, become tack-free in about 90
minutes, and become hard enough to file, sand,
drill and paint in 12 hours. They will stick to
almost everything except materials like Teflon,
polyethylene and waxed paper. They are resistant
to mild acids, caustics, detergents, gasoline and
fuel oil, fresh and salt water, and can be
applied to wet and submerged objects. Their
service temperature range is from -20 to 200�F

I have used them for many chores ranging from
patching an automobile fender to repairing the
handle of a frying pan, to sealing components of
a rough vacuum system together. I believe that
in the cured state they have very low vapor
pressures, but have no data on that matter. I
have used them successfully in rough vacuum
systems. I think they would probably be OK in
high vacuum applications (down to 10^-6 Torr),
and wouldn't be surprised if they would be OK in
the UHV range, but have no data on that. I have
been able to buy them at the local hardware
store. A package consisting of one 35 mm film
cartridge container of resin and one of hardener
usually costs less then $10.

Try 'em, you'll find them very useful.
--
Wilbur C. Bigelow, Professor Emeritus
Materials Sci. & Engr., Univ. of Michigan
Ann Arbor, Michigan 48109-2136
e-mail: bigelow-at-umich.edu;
Fx:734-763-4788; Ph:734-975-0858
Address mail to: 2911 Whittier Court
Ann Arbor, MI 48104-6731

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Dear colleagues,
I would like to draw to your attention that the postdoctoral position
below is still open. Please feel free to forward the announcement to
potentially interested people.
Regards
Beno�t Zuber

Workplace: Institute of Anatomy and Institute of Physiology, Medical
Faculty, University of Bern

Project Title: Structural study of the three-dimensional network of
fibroblasts and myofibroblasts in the working myocardium

Position: Postdoctoral position

Description: Fibroblasts form a three-dimensional cellular network
around cardiomyocytes in the intact myocardium. Under certain
pathological conditions, fibroblasts can differentiate to
myofibroblasts, which possess an intermediate phenotype between
fibroblasts and smooth muscle cells. Myofibroblasts can establish
electrical coupling with cardiomyocytes which may promote
arrhythmogenesis. This phenomenon has been recently studied in vitro.
However, it is unclear to which extent these findings can be
extrapolated to the in vivo situation. In particular, the
three-dimensional organization of the myofibroblast network and of the
sites of interaction between myofibroblasts and cardiomyocytes in vivo
are still ill-defined and need to be characterized in order to
understand their function. This question will be studied by the
successful candidate in collaboration with the groups of Beno�t Zuber
and Stephan Rohr. Using state of the art techniques in electron and
light microscopy, the candidate will conduct a comparative study of
the three dimensional architecture of fibroblasts and myofibroblasts
in well-controlled cell culture systems and in healthy and diseased
working myocardia from different species.

Requirements: The successful candidate holds a PhD, is highly
motivated to work on a challenging project and has good communication
skills. Demonstrated experience in electron microscopy is a
prerequisite.

Start: Upon mutual agreement.

Duration: Initially for 2 � 3 years

Application: Applications, including a letter of motivation, a CV, a
list of publications and 3 references should be sent by email.

Contact/Application Addresses: Prof. Beno�t Zuber
(benoit.zuber-at-ana.unibe.ch) or Prof. Stephan Rohr (rohr-at-pyl.unibe.ch)

Links to the institutions: Institute of Anatomy, Department of Physiology

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I have a question: How does the SEM-EDX decide whether As or Pb or both is present when it sees a substantial 10.5 kV peak?

Mohammed Yousuf
Senior Research Fellow
Qatar University
________________________________

رؤيتنا: أن تصبح جامعة قطر نموذجا للجامعة الوطنية في المنطقة، تتميز بنوعية التعليم والأبحاث، وبدورها الرائد في التنمية الاقتصادية والاجتماعية.

Our Vision: Qatar University shall be a model national university in the region, recognized for high quality education and research, and for being a leader of economic and social development.

==============================Original Headers==============================
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8, 41 -- From: Mohammed Yousuf {mdyousuf-at-qu.edu.qa}
8, 41 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
8, 41 -- Subject: SEM-EDX As or Pb analysis
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Hi Mohammed,

The quick answer is that the analyzer doesn't know how to distinguish
the peaks; it is up to the analyst using his brain.

The analyzer generally just does an energy match to the peaks, though
some of the manufacturers have improved the algorithms to reduce the
number of false IDs. Perhaps one of the manufacturers would like to
elaborate on whether they look for missing peaks, etc when doing element
IDs.

As a note how things used to be...An old analyzer I had quite frequently
told me I had Praseodymium in many of my samples when the sample
actually contained Chromium.

In this case, it should be simple to see by eye the difference between
Pb and As. For As, you would see only Ka and Kb peaks whereas Pb will
have a whole series of L lines. There will also be a difference in the
lower energy lines. The As L series are around 1 keV whereas Pb M series
is ~2keV where thy nicely overlap with S. Note that the higher energy
lines may be difficult to see in SEM; they are quite obvious in my TEM.

Cheers,
Henk

At 8/22/2012 4:47 AM, mdyousuf-at-qu.edu.qa wrote:
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} I have a question: How does the SEM-EDX decide whether As or Pb or both is present when it sees a substantial 10.5 kV peak?
}
} Mohammed Yousuf
} Senior Research Fellow
} Qatar University
} ________________________________
}
}
} رؤيتنا: أن تصبح جامعة قطر نموذجا للجامعة الوطنية في المنطقة، تتميز بنوعية التعليم والأبحاث، وبدورها الرائد في التنمية الاقتصادية والاجتماعية.
}
} Our Vision: Qatar University shall be a model national university in the region, recognized for high quality education and research, and for being a leader of economic and social development.
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} 8, 41 -- Subject: SEM-EDX As or Pb analysis
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

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12, 26 -- From: "Hendrik O. Colijn" {colijn.1-at-osu.edu}
12, 26 -- Subject: Re: [Microscopy] SEM-EDX As or Pb analysis
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Stephanie,
Paraformaldehyde alone should not permeabilize membranes, but you should investigate how the PF was made. Apparently if you boil the water while making PF you can also create contaminants (solvents like glycol), which is an alcohol.I would make a fresh batch careful not to go over 80C,and repeat your immunolabeling. If extraction occured during fixation, your cross-linked epitopes could
end up anywhere.

Michael Delannoy
Johns Hopkins SOM Microscope Facility.

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Dear Mohammed,
If you are exciting a 10.5 kV peak, you are also exciting the lower kV peaks of As and Pb. Look for these peaks. They are separate from each other, but there is the possibility that the lower kV peak overlap with other elements (like As L with Mg K). There is a predictable set of intensity ratios for every element's fingerprint. Use them to help eliminate possibilities.
Ken

On Aug 22, 2012, at 8:59 AM, colijn.1-at-osu.edu wrote:

}
}
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} Hi Mohammed,
}
} The quick answer is that the analyzer doesn't know how to distinguish
} the peaks; it is up to the analyst using his brain.
}
} The analyzer generally just does an energy match to the peaks, though
} some of the manufacturers have improved the algorithms to reduce the
} number of false IDs. Perhaps one of the manufacturers would like to
} elaborate on whether they look for missing peaks, etc when doing element
} IDs.
}
} As a note how things used to be...An old analyzer I had quite frequently
} told me I had Praseodymium in many of my samples when the sample
} actually contained Chromium.
}
} In this case, it should be simple to see by eye the difference between
} Pb and As. For As, you would see only Ka and Kb peaks whereas Pb will
} have a whole series of L lines. There will also be a difference in the
} lower energy lines. The As L series are around 1 keV whereas Pb M series
} is ~2keV where thy nicely overlap with S. Note that the higher energy
} lines may be difficult to see in SEM; they are quite obvious in my TEM.
}
} Cheers,
} Henk
}
}
}
} At 8/22/2012 4:47 AM, mdyousuf-at-qu.edu.qa wrote:
} }
} }
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} } I have a question: How does the SEM-EDX decide whether As or Pb or both is present when it sees a substantial 10.5 kV peak?
} }
} } Mohammed Yousuf
} } Senior Research Fellow
} } Qatar University
} } ________________________________
} }
} }
} } رؤيتنا: أن تصبح جامعة قطر نموذجا للجامعة الوطنية في المنطقة، تتميز بنوعية التعليم والأبحاث، وبدورها الرائد في التنمية الاقتصادية والاجتماعية.
} }
} } Our Vision: Qatar University shall be a model national university in the region, recognized for high quality education and research, and for being a leader of economic and social development.
} }
} }
} }
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}
} --
} Hendrik O. Colijn www.ceof.ohio-state.edu
} OSU Campus Electron Optics Facility colijn.1-at-osu.edu
} 040 Fontana Labs (614) 292-0674 (V)
} 116 W. 19th Ave. (614) 292-7523 (F)
} Columbus, OH 43210
}
} "Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."
}
}
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|_|"|_|"|_|_|"|_|"|_|"|_|_|"|_|"|_|_|"|_|"|_|"|_|_|
Kenneth JT Livi, PhD
Director, The High-Resolution Analytical Electron Microbeam Facility
of the Integrated Imaging Center
Departments of Earth and Planetary Sciences and Biology
Olin Hall
3400 N Charles Street
Johns Hopkins University
Baltimore, Maryland 21218 USA
| | | .| | .| | .| | .| | | :| | | .| | .| | .| | .| | | :|

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Listers

I am looking to replace (or repair) the LN2 dewar for a Gatan 600 Duomill.
This is the cylindrical dewar that sits on top-center of the 'work chamber'
and is used to improve the chamber vacuum. (Not the cold stage dewar that
sits below the work chamber.)

Gatan can no longer supply this part. Anyone have a retired Duomill with an
intact dewar they are willing to part with? We are very willing to negotiate
terms.

Alternatively, does anyone have a recommendation for a precision weld shop
that can make a cryo- and vacuum-tight weld on thin-walled stainless steel?

Cheers
Roger

Roger A. Ristau, PhD
Electron Microscopy Specialist
Institute of Materials Science
97 North Eagleville Road
University of Connecticut
Storrs, CT 06269
vox: 860-486-5453
fax: 860-486-4745
www.ims.uconn.edu/~micro/

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X-from: nicolas.brodusch-at-mcgill.ca ()

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Email: nicolas.brodusch-at-mcgill.ca
Name: Nicolas Brodusch

Organization: McGill University

Title-Subject: [Filtered] SEM/TEM - Biological sample preparation

Message: Hello all,

A customer brought us tissues for SEM/TEM observations. The samples were flash-frozen in isopentane
and immersed in OCT. As OCT is not suitable for electron microscopy under vacuum, do anyone has any
idea of how these samples can be used in SEM/TEM? Is there any possible exchange of OCT with a more
conventionnal fixative (glutaraldehyde, osmium tetroxide...). Any suggestions would be greatly
appreciated.
Many thanks.

Nicolas

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Dear all,
dear Nicolas,
sorry if I am long with this reply.

I wonder about the type of tissue your customer brought to you in a very {unusual manner} (guessing this customer did NOT speak to you BEFORE handed over his preps...(;-(( )

I would like to ask some questions (make some statements) just to clarify what your situation will be/ is:

ok, if I assume the tissue in OCT now would be {muscle} ...
you said: {flash-frozen in isopentane and [then?] immersed in OCT...

0-0: On site surgery: fresh tissue prepared, mounted in OCT - then flash frozen in isopentane [no buffer, no other cryoprotectant, no chemical fixation]
0-1: flash-frozen [only tissue?] on site in operating theatre (pre-cooled isopentane at nearly -50 to -80 degr. C, isopentane cooling down by LN2- to its freezing-point )
0-2: (immersion following in OCT at this stage implausible) stored at -80 degr. C ? or: stored at -20 to -25 degr C? (OCT mounting could be done)
0-3: customer handed over the deep frozen (at least -80 degr. C) / "normally frozen"(= stored at -25 degr C)
0-4: Question: actual storage, and storage conditions of frozen specimen?

only as suggestion(s):
1-1: use FS (Freeze Substitution)-Procedure (physical)
1-2: use FF (Freeze Fracture)-Procedure (physical)
1-3: (if you have access to an AFS-machine, perhaps this apparatus - depending on spec-size - could be involved in spec-prep)

Classical TEM- (or SEM-)Processing (chemical) - perhaps saving some ultrastructural details
1-2a: if stored in -80 degr C (spec in Al-foil(?): let warm up slowly until -25/-20 degr. C
1-2b: prepare fixatives (+ at least approx. 2% sucrose) as usual, cool down nearly to the lowest temperature without reaching freezing point (control, always swirling solutions)
1-2c: immerse the spec mounted in frozen OCT into cooled fixative (maintained at nearly the same or somehow lower temperature compared to OCT-clot)
1-2d: OCT compound will melt slowly, tissue will be fixed at the phase boundary bit by bit. Slowly increase temperature ( switching between storage for some time in the fridge, let stand at RT outside the fridge etc. To rotate or to swirl the specimen in vial by means of e.g. magnetic bead is of advantage)
1-2e: after full warming up to RT process as you would do routinely

[1-3: after dehydration to 70% CPD-processing for SEM ]

I have tried such (1-2a-1-2e) some years ago in two urgent cases of retrospective examination of (any preserved material left only in frozen state) muscle as well as skin (I don't have FF, FS or CPD-possibility in our Lab).
It was possible to achieve an acceptable "ultrastructural" preservation, at least at a small border zone for TEM and really "good" (= not excellent but sufficient) for semithin section morphology. Quality of ultrastructural preservation also my be dependent on storage conditions (i. e. how many unforeseen "recrystallization cycles" the tissue [had to] underwent/was subjected to). Perhaps some practical alteration(s) in the procedure outlined here would also be helpful. If you try, would you let us know about the outcome?

Knowing the following substance to be really hazardous, namely ACROLEIN ( {propenal} , simplest unsaturated aldehyde) it might be interesting that its freezing point = minus 125 degr. F = -87 degr. C = 186 degr. K , so a mixture of (buffered?) Acrolein-FA-GA (acrolein is very fast in fixing tissue) perhaps would be advantageous..

Dear Nicolas:
Best wishes , good luck, and
looking forward to perhaps your sending all the offlist-replies you'll get in reply to your question, really an interesting task, I think..

Wolfgang MUSS
Member of MSA
SALZBURG-AUSTRIA

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Hello all,
A customer brought us tissues for SEM/TEM observations.
The samples were flash-frozen in isopentane and immersed in OCT.
As OCT is not suitable for electron microscopy under vacuum, do anyone has any idea of how these samples can be used in SEM/TEM?

Is there any possible exchange of OCT with a more conventionnal fixative (glutaraldehyde, osmium tetroxide...).
Any suggestions would be greatly
appreciated.
Many thanks.

Nicolas

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Title-Subject: [Filtered] TEM Method for Algae

Message: Hello All.

I have a scientist that will be bring Algae samples for TEM thin sectioning. I have not prepped
this kind of sample before. Does anyone have a method for the dehydration and embedding?

Thanks,
Michelle

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Dear Nicolas

The diagnostic pathology lab where I work routinely uses a similar technique for freezing muscle tissue. The tissue is mounted in OCT and then plunged into the isopentane which has been frozen to a slurry by holding the beaker in liquid nitrogen. Alternatively OCT tissue is frozen directly in liquid nitrogen for routine frozen sectioning.

OCT is water soluble so tissue is transferred directly to 10% neutral buffered formalin for routine histological processing. If TEM was required it would be transferred to glutaraldehyde.

If the specimens have been stored at -80 degC then transferring to -20 degC as suggested by Wolfgang would be preferred to prevent the tissue cracking due to sudden temperature change. Then to fixative at RT. I understand that any freezing artefact has already occurred at the time of freezing or during storage.

An important point to remember is to remove the OCT completely to achieve adequate fixation. Swirling the jar is effective. And as I found out the hard way in my junior years, OCT forms a hard white solid when it comes into contact with alcohol. A chemist amongst us will no doubt enlighten us on this reaction between the alcohol and the resin component of OCT!!

Hope this helps
Naomi

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Title-Subject: [Filtered] SEM/TEM - Biological sample preparation

Message: Hello all,

A customer brought us tissues for SEM/TEM observations. The samples were flash-frozen in isopentane
and immersed in OCT. As OCT is not suitable for electron microscopy under vacuum, do anyone has any
idea of how these samples can be used in SEM/TEM? Is there any possible exchange of OCT with a more
conventionnal fixative (glutaraldehyde, osmium tetroxide...). Any suggestions would be greatly
appreciated.
Many thanks.

Nicolas

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Two of MSA's educational resources are now easier to use! Go to the
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Opportunities" menu. If you go to "Project MICRO" (MSA's middle
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Email: mamiller-at-coh.org
Name: Marcia Miller

Organization: Beckman Research Institute, City of Hope

Title-Subject: [Filtered] How to do TEM on 2000 cells?

Message: Dear Electron Microscopists,

We have a user of our EM Core who would like TEM images from a population of cells collected by flow
sorting. The cell number is really tiny, just around 2000 cells. The best I can come up with is to
pellet the cells, noting where the cells land (by example in another tube) on the inside of the tube
(1.5 ml), then after removing the medium without disturbing the pellet of 2000 cells, add an aliquot
of non-relevant cells, such as RBCs, to augment the pellet to the point of visibilitiy, and pellet
again being sure that the added cells will land on the same area as the 2000. I invision the
precious 2000 being cupped by the cells from the augmenting aliquot. Does this sound reasonable?
Has anyone done something like this? Is there any method that can be used with confidence? The
cells are neurons from specific mounse ganglia and the objective is to look for herpesviruses
inside. We will be happy to receive any comments or suggestions. Thanks, Marcia Miller, EM Core,
Beckman Research Institute, City of Hope, Duarte CA 91010

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Marcia,
Have you considered using agarose to gel the tiny pellet after fixation?

sincerely;
Michael Delannoy

----- Original Message -----
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Hello Marcia,

Vlad Speransky (NIH) had a similar question in May this year and the possibility to use microdialysis tubing as is used in high pressure freezing applications emerged (Hohenberg, H., Mannweiler, K., and Mueller, M. (1994). High-pressure freezing of cell suspensions in cellulose capillary tubes. J. Microsc. 175, 34�43). Cells might be collected in there (in fixative perhaps) and the tubing clamped shut. That way you would have a tiny packet that does not get lost as easily.

Good luck!

Jan Leunissen
Dunedin, New Zealand
i: www.aurion.nl

On 25/08/2012, at 12:14 AM, microscopylistserver-noreply-at-microscopy.com wrote:

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}
} We have a user of our EM Core who would like TEM images from a population of cells collected by flow
} sorting. The cell number is really tiny, just around 2000 cells. The best I can come up with is to
} pellet the cells, noting where the cells land (by example in another tube) on the inside of the tube
} (1.5 ml), then after removing the medium without disturbing the pellet of 2000 cells, add an aliquot
} of non-relevant cells, such as RBCs, to augment the pellet to the point of visibilitiy, and pellet
} again being sure that the added cells will land on the same area as the 2000. I invision the
} precious 2000 being cupped by the cells from the augmenting aliquot. Does this sound reasonable?
} Has anyone done something like this? Is there any method that can be used with confidence? The
} cells are neurons from specific mounse ganglia and the objective is to look for herpesviruses
} inside. We will be happy to receive any comments or suggestions. Thanks, Marcia Miller, EM Core,
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Name: Barbara Plowman

Organization: University of the Pacific Arthur A. Dugoni School of Dentistry

Title-Subject: [Filtered] EM on Flow cytometry

Message: Yes, I was a student guest in a Lawrence Livermore lab in 1977 where they had a flow
cytometer, even before they were commercial. We did TEM on some collected cells but did not pellet
them in agar. I was told , " they looked a little beat up"...

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Title-Subject: [Filtered] Question about Searchable Image Database

Message: Good Day Everyone:
I have enjoyed reading and replying to posts for many, many years now and find the expertise and
advice exceptional. I have a quick question for the group. We have been developing an image
repository for several years now, and we want to make an image database that is searchable, secure,
archived, and somewhat user friendly. We are going to ensure that specific metadata from images are
in searchable fields. Metadata we will capture include voltage, scale bar information, initial
magnification, time stamp, name, etc. The software will have to be from a US vendor (if possible)
and will have to pass US government security settings. This will ultimately be a secure searchable
database that has the ability to be expanded in the future. The software will be hosted on our own
secure servers, and will not be linked to the internet. The initial database will contain
approximately 5,000 images. Any and all replies are most welcome.

Thank you in advance,

Robert

Dr. Robert K. Pope

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Dear Marcia,

There are many approaches for preparing small numbers of cells for TEM examination (flat embedding them on glass, centrifuging them to a specific location in a tube, pelleting them down in a sealed Eppendorf pipette tip, embedding them in agarose or gelatin etc). You could contact me off-line so we could discuss options and find a good approach to use.

Your real problem will be in preserving the morphology of cells that have been through a cell sorter. It is a really disruptive process for the cells to go through and considering what they look like in the TEM, it is surprising that they are still viable. Is there a way for the researcher to let the cells recover before fixation? If not, then you will have to use the material given to you. Centrifugation is a problem because the cells will be very fragile, but there are ways around that too.

Regards,

Paul.

Paul Webster, Ph.D.
House Research Institute
2100 West Third Street
Los Angeles
CA 90057.

-----Original Message-----
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Sent: Fri 8/24/2012 5:39 AM
To: Webster, Paul

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Name: Marcia Miller

Organization: Beckman Research Institute, City of Hope

Title-Subject: [Filtered] How to do TEM on 2000 cells?

Message: Dear Electron Microscopists,

We have a user of our EM Core who would like TEM images from a population of cells collected by flow
sorting. The cell number is really tiny, just around 2000 cells. The best I can come up with is to
pellet the cells, noting where the cells land (by example in another tube) on the inside of the tube
(1.5 ml), then after removing the medium without disturbing the pellet of 2000 cells, add an aliquot
of non-relevant cells, such as RBCs, to augment the pellet to the point of visibilitiy, and pellet
again being sure that the added cells will land on the same area as the 2000. I invision the
precious 2000 being cupped by the cells from the augmenting aliquot. Does this sound reasonable?
Has anyone done something like this? Is there any method that can be used with confidence? The
cells are neurons from specific mounse ganglia and the objective is to look for herpesviruses
inside. We will be happy to receive any comments or suggestions. Thanks, Marcia Miller, EM Core,
Beckman Research Institute, City of Hope, Duarte CA 91010

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Colleagues,

I am searching for *defective* image acquisition and bus expansion
boards from Micrion and FEI-Vectra Focused Ion Beam tools.

These parts were considered "unrepairable" by the OEM and typically were
either abandoned at customer's lab or discarded to trash. I can't offer
to pay for the defective PCBs, but would be more then happy to pay
shipping charges if someone has one or more collecting dust in a junk
drawer under the bench. The PCBs that I am interested to collect are:

1. "Piranha" image acquisition board for MCA bus. It may have UDC8032
and UPX8400 part numbers on it, was made by former Univision
Technologies and installed on Micrion FIB tools with PowerServer 370
Power PC.

2. "Falcon PCI" image acquisition board for PCI bus. It may have 199678
part number on it, was made by former Univision Technologies Company and
later for some period of time produced by Titan Corporation. This board
was installed in Micrion and later FEI "Vectra" series of FIB systems
with p43 Power PC.

3. "MCA to Multibus" and "PCI to Multibus" adapter boards, models
444-201 and 444-202. These boards were made by former Bit3 Computer
Corporation and may have 82801120, or 82402025, or other part numbers,
and were installed in Micrion and later FEI "Vectra" series of FIB
systems with both Powerserver 370 and p43 Power PC.

I am mostly interested in non-functioning boards without physical
damage, but would take board in any condition.

Thank you very much beforehand,
--
Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
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Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
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Dear Marcia,

My first thought when I read your message was the same as Paul: if the user wants to use TEM, it most probably means that he wants a good morphology.
Of course the previous propositions are all valid techniques, but you may just be able to optimally prepare damage/unhealthy cells and then I wonder what sense that makes.
If the cells are still viable after cell sorting, you may put them in a sterile petri dish with some medium and let them adhere to�a suitable substrate for flat embedding. In this case 2000 cells are more then enough. Most cells adhere within several hours, this would leave them some time to recover too.
May I advise you to consider taking appropriate measures when working� with infected material (including disposal), especially when the material is infected with a human virus?
�
Best regards,
Stephane

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Cc:
Sent: Sunday, August 26, 2012 9:09 PM

Dear Marcia,

There are many approaches for preparing small numbers of cells for TEM examination (flat embedding them on glass, centrifuging them to a specific location in a tube, pelleting them down in a sealed Eppendorf pipette tip, embedding them in agarose or gelatin etc). You could contact me off-line so we could discuss options and find a good approach to use.

Your real problem will be in preserving the morphology of cells that have been through a cell sorter. It is a really disruptive process for the cells to go through and considering what they look like in the TEM, it is surprising that they are still viable. Is there a way for the researcher to let the cells recover before fixation? If not, then you will have to use the material given to you. Centrifugation is a problem because the cells will be very fragile, but there are ways around that too.

Regards,

Paul.

Paul Webster, Ph.D.
House Research Institute
2100 West Third Street
Los Angeles
CA 90057.

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Email: mamiller-at-coh.org
Name: Marcia Miller

Organization: Beckman Research Institute, City of Hope

Title-Subject: [Filtered] How to do TEM on 2000 cells?

Message: Dear Electron Microscopists,

We have a user of our EM Core who would like TEM images from a population of cells collected by flow
sorting.� The cell number is really tiny, just around 2000 cells.� The best I can come up with is to
pellet the cells, noting where the cells land (by example in another tube) on the inside of the tube
(1.5 ml), then after removing the medium without disturbing the pellet of 2000 cells, add an aliquot
of non-relevant cells, such as RBCs, to augment the pellet to the point of visibilitiy, and pellet
again being sure that the added cells will land on the same area as the 2000.� I invision the
precious 2000 being cupped by the cells from the augmenting aliquot.� Does this sound reasonable?
Has anyone done something like this?� Is there any method that can be used with confidence?� The
cells are neurons from specific mounse ganglia and the objective is to look for herpesviruses
inside. We will be happy to receive any comments or suggestions. Thanks, Marcia Miller, EM Core,
Beckman Research Institute, City of Hope, Duarte CA 91010

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Title-Subject: [Filtered] Job vacancy: Image Analysis and Computed Tomography Officer at the
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Message: Dear all,

The University of Sydney is currently recruiting for the following position.

IMAGE ANALYSIS AND COMPUTED TOMOGRAPHY OFFICER
SYDNEY MICROSCOPY AND MICROANALYSIS (SMM)

Image analysis experience essential
Previous experience in an educational environment will be highly regarded
Full-time continuing, remuneration package: $92K - $100K p.a. which includes leave loading and up
to 17% super

The mission of the SMM is to provide a core set of state-of-the-art microscopy and microanalysis
services to researchers.

We are currently recruiting an Image Analysis and Computed Tomography Officer to coordinate user
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This is a unique opportunity to further develop your professional career and join one of Australias
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All applications must be submitted via the University of Sydney careers website. Visit
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Title-Subject: [Filtered] Freeze Dryer

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I am looking for a freeze drying device for sample preparation for conventional scanning electron
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I'm sorry but this is the wrong forum Valery

On 8/26/2012 7:32 PM, vray-at-partbeamsystech.com wrote:
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} Colleagues,
}
} I am searching for *defective* image acquisition and bus expansion
} boards from Micrion and FEI-Vectra Focused Ion Beam tools.
}
} These parts were considered "unrepairable" by the OEM and typically were
} either abandoned at customer's lab or discarded to trash. I can't offer
} to pay for the defective PCBs, but would be more then happy to pay
} shipping charges if someone has one or more collecting dust in a junk
} drawer under the bench. The PCBs that I am interested to collect are:
}
} 1. "Piranha" image acquisition board for MCA bus. It may have UDC8032
} and UPX8400 part numbers on it, was made by former Univision
} Technologies and installed on Micrion FIB tools with PowerServer 370
} Power PC.
}
} 2. "Falcon PCI" image acquisition board for PCI bus. It may have 199678
} part number on it, was made by former Univision Technologies Company and
} later for some period of time produced by Titan Corporation. This board
} was installed in Micrion and later FEI "Vectra" series of FIB systems
} with p43 Power PC.
}
} 3. "MCA to Multibus" and "PCI to Multibus" adapter boards, models
} 444-201 and 444-202. These boards were made by former Bit3 Computer
} Corporation and may have 82801120, or 82402025, or other part numbers,
} and were installed in Micrion and later FEI "Vectra" series of FIB
} systems with both Powerserver 370 and p43 Power PC.
}
} I am mostly interested in non-functioning boards without physical
} damage, but would take board in any condition.
}
} Thank you very much beforehand,

--
Robert Keyse
EM Facility
Whitaker Laboratory
5 East Packer Avenue
Bethlehem
PA 18015
USA

Tel. +1 610 758 4283
Fax +1 610 758 4244

==============================Original Headers==============================
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I am investigating fungal-plant host-parasite interactions. So, I do
not want to use critical point drying and put the samples in fixation
solution because this would wash away fungal exudates and oxalate
crystals. Therefore, I think freeze drying would be the best method
for preparation.

Anne

Zitat von "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu} :

} I'm not a biologist, so I am making a stretch here.
}
} I know drying is necessary to bad things don't happen to the sample
} and microstructure in the SEM.
}
} I'd be interested to know why your sample cannot be exposed to
} liquids. I suppose the biologists that got your note would also want
} to know. That restriction may really limit your options.
}
} I thought the idea of critical point drying was to substitute ne
} liquid for another and then remove it by going around critical point
} to avoid the gas-liquid interface and surface tensions and the havoc
} they would wreak on the sample.
}
} I also understand that flash freezing is sometimes used followed by
} freeze-drying. Apparently the solid-gas interface is not so
} problematic. The freezing must be done very quickly so that the
} solid-liquid transition does not change the structure. The ice
} crystals must be kept small or even non-existent by forming
} amorphous ice. That probably requires the samples to be very small
} or thin.
}
} I understand such devices are available. Of course the question
} remains as to whether your samples are amenable to the process. I
} seem to recall that it involves plunging the sample into some kind
} of cold slush.
}
} Warren
}
} -----Original Message-----
} From: microscopylistserver-noreply-at-microscopy.com
} [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: Monday, August 27, 2012 7:23 AM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] viaWWW:Freeze Dryer
}
}
}
}
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} X-from: Anne.Heller-at-uni-hohenheim.de ()
}
} This Question/Comment was submitted to the Microscopy Listserver
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} Email: Anne.Heller-at-uni-hohenheim.de
} Name: Anne Heller
}
} Organization: Institut f�r Botanik, Universit�t Hohenheim
}
} Title-Subject: [Filtered] Freeze Dryer
}
} Message: Dear All,
}
} I am looking for a freeze drying device for sample preparation for
} conventional scanning electron
} microscopy not cryo scanning electron microscopy. We cannot use
} critical point drying, because the
} samples should not come in contact with liquids. The samples will be
} frozen, freeze dried,
} sputtered, and investigated in a conventionel SEM. Are there any
} practicable devices on the market?
}
} Anne Heller
}
} Login Host: 144.41.33.113
} Listserver Email Form V - 20120416
} ---------------------------------------------------------------------------
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}
}
}
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} ==============================Original Headers==============================
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}

Dr. Anne Heller
AG Elektronenmikroskopie
Institut f�r Botanik (210)
Universit�t Hohenheim
Postfach, 70593 Stuttgart
Tel. 0711-45922180
Fax. 0711-45923355

==============================Original Headers==============================
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Anne,

Have you considered doing osmium vapor fixation? You can put your sample in a closed container with exposed osmium fixative (fume hood, gloves!) for some period, then afterwards mount it onto a viewing stub for carbon or metal coating and view like that. It is sometimes possible to view specimens that have not been completely dehydrated, but normally that is best left for a variable pressure/environmental SEM. Of course, if you have one of these, you can often avoid any fixation whatsoever.

Good luck,
Randy

Randy Tindall
Senior Research Specialist
Electron Microscopy Core Facility
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65201
(573) 884-5383

We Do Small Well!

www.emc.missouri.edu (dayjob)
http://nadiasyard.com (blog)
http://www.facebook.com/RandyTindallGalleries (photography)

-----Original Message-----
X-from: Anne.Heller-at-uni-hohenheim.de [mailto:Anne.Heller-at-uni-hohenheim.de]
Sent: Monday, August 27, 2012 10:30 AM
To: Tindall, Randall D.

I am investigating fungal-plant host-parasite interactions. So, I do not want to use critical point drying and put the samples in fixation solution because this would wash away fungal exudates and oxalate crystals. Therefore, I think freeze drying would be the best method for preparation.

Anne

Zitat von "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu} :

} I'm not a biologist, so I am making a stretch here.
}
} I know drying is necessary to bad things don't happen to the sample
} and microstructure in the SEM.
}
} I'd be interested to know why your sample cannot be exposed to
} liquids. I suppose the biologists that got your note would also want
} to know. That restriction may really limit your options.
}
} I thought the idea of critical point drying was to substitute ne
} liquid for another and then remove it by going around critical point
} to avoid the gas-liquid interface and surface tensions and the havoc
} they would wreak on the sample.
}
} I also understand that flash freezing is sometimes used followed by
} freeze-drying. Apparently the solid-gas interface is not so
} problematic. The freezing must be done very quickly so that the
} solid-liquid transition does not change the structure. The ice
} crystals must be kept small or even non-existent by forming amorphous
} ice. That probably requires the samples to be very small or thin.
}
} I understand such devices are available. Of course the question
} remains as to whether your samples are amenable to the process. I seem
} to recall that it involves plunging the sample into some kind of cold
} slush.
}
} Warren
}
} -----Original Message-----
} From: microscopylistserver-noreply-at-microscopy.com
} [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: Monday, August 27, 2012 7:23 AM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] viaWWW:Freeze Dryer
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
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} X-from: Anne.Heller-at-uni-hohenheim.de ()
}
} This Question/Comment was submitted to the Microscopy Listserver
} using the WWW based Form at
} http://microscopy.com/MicroscopyListserver/MLFormMail.html
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} Microscopy Listserver
} ---------------------------------------------------------------------------
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} Email: Anne.Heller-at-uni-hohenheim.de
} Name: Anne Heller
}
} Organization: Institut f�r Botanik, Universit�t Hohenheim
}
} Title-Subject: [Filtered] Freeze Dryer
}
} Message: Dear All,
}
} I am looking for a freeze drying device for sample preparation for
} conventional scanning electron
} microscopy not cryo scanning electron microscopy. We cannot use
} critical point drying, because the
} samples should not come in contact with liquids. The samples will be
} frozen, freeze dried,
} sputtered, and investigated in a conventionel SEM. Are there any
} practicable devices on the market?
}
} Anne Heller
}
} Login Host: 144.41.33.113
} Listserver Email Form V - 20120416
} ---------------------------------------------------------------------------
}
}
}
}
} ---===[|]===---
}
}
}
} --
} ===========================================
} Do not reply to this message it is from
} the Microscopy Listserver NO-REPLY forwarding
} system. You should send a new message to
}
} Microscopy-at-Microscopy.com
}
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}
} ==============================Original Headers==============================
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} 27 07:22:11 2012
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Dr. Anne Heller
AG Elektronenmikroskopie
Institut f�r Botanik (210)
Universit�t Hohenheim
Postfach, 70593 Stuttgart
Tel. 0711-45922180
Fax. 0711-45923355

==============================Original Headers==============================
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9, 37 -- Date: Mon, 27 Aug 2012 17:19:08 +0200
9, 37 -- From: "Dr. Anne Heller" {Anne.Heller-at-uni-hohenheim.de}
9, 37 -- To: Microscopy-at-microscopy.com
9, 37 -- Subject: RE: [Microscopy] viaWWW:Freeze Dryer
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Dear Anne,

we bought a freeze dryer recently that should be suitable for SEM . For
further details or if you want to test it please contact me off the list
server.

Regards,
Baerbel

--
Dr. Baerbel Hauroeder
ZInstSanBw Koblenz
Elektronenmikroskopie
Andernacherstr. 100
56070 Koblenz
Tel.: +49 261-896 7260
Fax: +49 261-896 7269
Mail:b.hauroeder(at)zinstkob.de

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Dear Listserver,

Is there anyone who can provide advice/instructions/warnings, etc. with
respect to proper (legal) disposal of uranyl acetate in the U.S.?

Thanks, Bill Roberts

DuPont Teijin Films
William.H.Roberts-at-usa.dupont.com

This communication is for use by the intended recipient and contains
information that may be Privileged, confidential or copyrighted under
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Dear colleagues,
�
I am a biologist and I am a little bit lost with a mineralogist issue.
Someone asked me to quantify mineral particles in a powder (number of particles�per mg powder). The particles have a size distribution compatible with LM (0,4-10�m).
I could just weight a small amount of powder (5 mg) on a glass slide and count the particles in LM but there are too many particles and weighting less than 5 mg with precision is not realistic.
So alternatively I could suspend the particles and put a certain volume on a glass slide, dry the liquid and count the particles on a LM but I fear that the particles would agglomerate.
Is it realistic to think that I could resuspend the particles in say methanol or acetone, sonicate the suspension and quickly put (quantitatively) a small amount on a glass slide (eventually on a heating plate)?
I could probably do something similar with a stub and SEM, but this would probably unnecessarily increase the analysis time (however I could clearly see agglomerates).

I would appreciate it if you'd share your experience and thoughts with me. Even the weirdest ones (which sometimes turn out to be the best).
Any other method will be considered too, as far as I can practically use it. I also have�access to a TEM and a SEM.

Many thanks in advance.

Stephane

==============================Original Headers==============================
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5, 38 -- Reply-To: Stephane Nizet {nizets2-at-yahoo.com}
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Dear Erin,

I believe the jet will work well even if you change the position of the
Y shaped part inside the assembly. But the jet bottom should touch the
bottom of the pump otherwise the top of the jet will be too hight. The
jet assembly for 4" diffusion pump is the same for all JEOL microscope
with diffusion pump. I hope you can get one on an old machine somewhere
in your country. I'm interesting to know what kind of contamination was
in the pump and how it came in.
Regards

--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

microscopylistserver-noreply-at-microscopy.com a �crit :
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} Email: ewood1-at-uvm.edu
} Name: Erin Wood
}
} Organization: University of Vermont-School of Engineering
}
} Title-Subject: [Filtered] JEOL JSM-6060 SEM jet assembly
}
} Message: Hello everyone,
}
} I hope this inquiry reaches you all in good health and spirits! I am a graduate student in the
} mechanical engineering program at the University of Vermont.
} I am curious if anyone has any experience with JEOL diffusion pumps, specifically, I believe the 4
} inch pump. It seems that our diffusion pump maybe had some contamination in it so we tried to take
} it apart to clean, and the aluminium shoulder screws at the bottom got stuck in the little Y shaped
} connector in the jet assembly. Long story short, we are at the point where we will now try to drill
} out the Y-shaped connector and retap it, and get some new shoulder screws. If this fails we are
} looking for a back up plan, so I thought as a last ditch effort we would see if anyone had this
} piece from a jet assembly that perhaps is no longer able to be used. This would be easier than
} buying the entire jet assembly, as ours is reconditioned and ready to go we think.
}
} We are also curious if the assembly is sitting slightly lower or higher due to the new screws and
} threading, will this affect our pumping at all? I'm not sure that contamination was actually the
} problem, but it is our best guess that it was. But I'd hate to put the pump back together and have
} it not work because it was sitting slightly off from the original position.
}
} Thank you so much for reading this, and for any advice you may offer.
}
} Erin L Wood
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9, 25 -- From Nicolas.Stephant-at-univ-nantes.fr Wed Aug 29 02:29:41 2012
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Dear All,

I am new to the list, so I hope this type of email is not of topic.

I am an assistant professor at the Delft Technical University in the
Netherlands. I am very interested in applying principles of control
theory to automate TEMs. I am currently developing a project proposal
for the European Research Council (ERC) for which I would like to find
partners.

Specifically, I am looking for a young expert (with a Ph.D. of less
than 6 years) in electron optics modeling and simulation (i.e.,
simulation of electron trajectories and magnetic fields inside a TEM).
He/she would have to be located in either Europe (except The
Netherlands), its associated countries, the BRIC countries or Latin
America.

If you know of anyone that could be interested, please email me
directly (not to bother the list).

Thank you for your help.

Regards,

Arturo Tejada

===================================================
Dr. Arturo Tejada
Assistant Professor
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Email: nicolas.brodusch-at-mcgill.ca
Name: Nicolas Brodusch

Organization: McGill University, Montr�al

Title-Subject: [Filtered] SEM/TEM - Biological sample preparation

Message: Hello all,

Thanks to all of you who answered my questions with useful information. Concerning the way specimens
where frozen this is all the customer sent to me:"It seems as though all they use is an isopentane
dry ice bath to flash freeze the tissue, and then place the specimen in OCT". For the moment,
samples are stored at -80C. The customer would like to image the mitochondria's structure in thyroid
tissus. Do you think that imaging mitochondria's structure would be possible if I'm able to remove OCT?

Many thank

Best regards,

Nicolas

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Just a thought off the top of my head. I would want to know what the material is to insure it doesn't dissolve in my mounting media. I'd want the refractive index of the particles sufficiently different from the mounting media and I would "suspend" a known weight in a volume of water and count then like red blood cells. I would not ultrasonicate for fear of making agglomerates in to smaller particles.

If that didn't work , I try the same approach, suspend in a more viscous, low evaporation rate media (I'm measuring metal particles in immersion oil right now), transfer a small amount to a slide add a sufficiently large cover glass. I'd use a mechanical stage to sweep the entire area and count them.

Being old fashion, I would use the mark 1 eyeball and a tally counter and would try to limit the population to 100-200 particles per cover glass.

Knowing all your volumes and weights, I'd back calculate the results.

Sounds interesting. Let me how it turns out and how you did it.

Stay safe...........
Frank

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Wednesday, August 29, 2012 3:41 AM
To: Frank Karl

Dear colleagues,

I am a biologist and I am a little bit lost with a mineralogist issue.
Someone asked me to quantify mineral particles in a powder (number of particles per mg powder). The particles have a size distribution compatible with LM (0,4-10�m).
I could just weight a small amount of powder (5 mg) on a glass slide and count the particles in LM but there are too many particles and weighting less than 5 mg with precision is not realistic.
So alternatively I could suspend the particles and put a certain volume on a glass slide, dry the liquid and count the particles on a LM but I fear that the particles would agglomerate.
Is it realistic to think that I could resuspend the particles in say methanol or acetone, sonicate the suspension and quickly put (quantitatively) a small amount on a glass slide (eventually on a heating plate)?
I could probably do something similar with a stub and SEM, but this would probably unnecessarily increase the analysis time (however I could clearly see agglomerates).

I would appreciate it if you'd share your experience and thoughts with me. Even the weirdest ones (which sometimes turn out to be the best).
Any other method will be considered too, as far as I can practically use it. I also have access to a TEM and a SEM.

Many thanks in advance.

Stephane

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Many thanks for�helping.
Most of you�proposed using an hemacytometer to count the particles.
It is my (limited) understanding that mineral particles around 1�m while stay in suspension for a long time, probably more than 24h. In this condition it will be impossible to count the particles because they'll swim all the time.
The solution must take the particle size into account, it is not a trivial question. Particles above 10�m will definitely sediment within minutes, do not agglomerate and are easy to filter, but the problem is more complex for small particles.
Considering a SEM analysis, I wonder how I could reconcile the small field of view with the necessity to count quantitatively (completely) all the particles in the stub. How could I know which particle I have already counted and not count it twice?

I would love to have access to a particle counter�but this is not the case.

Stephane with a smoking brain

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Hi Stephane,
Why not taking images of your fields and count them from the image, by hand, or with ImageJ ;-)
You restrict sedimentation, and the possibility of counting particles twice, if you can collect your fields fast enough.

Joachim

Seventhwave Laboratories
St. Louis

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Wednesday, August 29, 2012 8:34 AM
To: Joachim Siegmund

Many thanks for�helping.
Most of you�proposed using an hemacytometer to count the particles.
It is my (limited) understanding that mineral particles around 1�m while stay in suspension for a long time, probably more than 24h. In this condition it will be impossible to count the particles because they'll swim all the time.
The solution must take the particle size into account, it is not a trivial question. Particles above 10�m will definitely sediment within minutes, do not agglomerate and are easy to filter, but the problem is more complex for small particles.
Considering a SEM analysis, I wonder how I could reconcile the small field of view with the necessity to count quantitatively (completely) all the particles in the stub. How could I know which particle I have already counted and not count it twice?

I would love to have access to a particle counter�but this is not the case.

Stephane with a smoking brain

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"When you are a hammer, every problem looks like a nail."

Being an SEM-EDS guy, I do think of using SEM rather than light microscopy. I think the analysis could be done rather quickly and might render it more cost-effective than LM.

You have not said what the two phases are - the mineral and the powder. If they are significantly different, I would consider using EDS to determine their relative abundance, SEM to determine the average particle size, and then use known densities to calculate the number of particles in a volume (mass) of sample.

Of course that assumes that there is some chemical differentiation between mineral and powder. It also would help if the powder is not an organic material. It would be hard to do a decent chemical analysis in that case. If that were the case, it might be easier to determine the mass loading by some other chemical analysis and use the average particle size from SEM to go through the calculation.

Warren

-----Original Message-----
X-from: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
Sent: Wednesday, August 29, 2012 2:29 AM
To: wesaia-at-iastate.edu

Dear colleagues,
�
I am a biologist and I am a little bit lost with a mineralogist issue.
Someone asked me to quantify mineral particles in a powder (number of particles�per mg powder). The particles have a size distribution compatible with LM (0,4-10�m).
I could just weight a small amount of powder (5 mg) on a glass slide and count the particles in LM but there are too many particles and weighting less than 5 mg with precision is not realistic.
So alternatively I could suspend the particles and put a certain volume on a glass slide, dry the liquid and count the particles on a LM but I fear that the particles would agglomerate.
Is it realistic to think that I could resuspend the particles in say methanol or acetone, sonicate the suspension and quickly put (quantitatively) a small amount on a glass slide (eventually on a heating plate)?
I could probably do something similar with a stub and SEM, but this would probably unnecessarily increase the analysis time (however I could clearly see agglomerates).

I would appreciate it if you'd share your experience and thoughts with me. Even the weirdest ones (which sometimes turn out to be the best).
Any other method will be considered too, as far as I can practically use it. I also have�access to a TEM and a SEM.

Many thanks in advance.

Stephane

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} realname - Jamie Carr
} Email - jamie.carr-at-state.ma.us
} ORGANIZATION - MA DCR
} EDUCATION - Graduate College
} LOCATION - MA
} SUBJECT_OF_QUESTION - Microscope camera
} QUESTION - Hello,
} My name is Jamie Carr and I am an Aquatic Biologist in
} Massachusetts. A colleague suggested that I post a question to the
} "Ask a Microscopist" listserve.
} I am in the process of purchasing a new inverted compound Olympus
} microscope (probably a BX43) for our department to fulfill our role
} of identifying and enumerating algae and phytoplankton in a
} reservoir to ensure drinking water quality. We currently have an
} older Reichert scope and no imaging capability. We are typically
} working at magnifications from 100 to 200x. Our budget is 10-12K.
} It seems that Olympus typically packages their scopes with
} Lumenera digital cameras, but it seems to me that many well
} respected labs use the Zeiss Axiocam. I am rather frustrated by the
} lack of reviews and independent information available online that
} might help me compare cameras. I am working to try and be able to
} demo the Olympus scope with a Lumenera camera (~Infinity 2, 3
} megapixel model) and possibly a Zeiss (~MRc, 2 megapixel model) as
} well.
} I have tried to research this as much as possible, but I still
} feel a bit under informed and would appreciate any commentary based
} on experience. Are these two cameras both quality choices? Should
} I be considering any other cameras to pair with this scope? It's a
} bit confusing to me as more megapixels are not always better, but is
} it worth spending a lot more $ for more megapixels or better
} quality/name with fewer megapixels?
} Any general input or suggestions would be greatly appreciated..
} Thanks, Jamie

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Email: derrick.horne-at-botany.ubc.ca
Name: Derrick Horne

Organization: UBC BioImaging Facility

Title-Subject: [Filtered] Volume increase post-CPD:references?

Message: I know we all talk about samples being hygroscopic after CPD, but I am having a difficult
time coming up with any references where investigators have looked at volume change in CPD tissue
due to water absorption, post-CPD.

I have a user who is going to be doing MicroCT on critically point dried lung cores (10mmx20mm), and
the cores will not be post-fixed in osmium tetroxide for various reasons.

Each CT could take up to 3h. The question the researcher has is whether or not there will be
appreciable volume change over this time in a core, stored sealed from air post-CPD, that is then
exposed to air immediately prior to being mounted and scanned over 3h.

Is anyone aware of anything in the literature that addresses this?

Thanks in advance

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On Aug 28, 2012, at 1:19 PM, William.H.Roberts-at-usa.dupont.com wrote:

} Dear Listserver,
}
} Is there anyone who can provide advice/instructions/warnings, etc.
} with
} respect to proper (legal) disposal of uranyl acetate in the U.S.?
}
} Thanks, Bill Roberts

Dear Bill,
Since no one else has tackled this, I'll give it a shot. My info is
more than a decade old; however, at least this part is still valid:
Different states have different rules, so the correct answer will
depend on where you plan to dispose of the UAc. When I was last
confronted with this issue, I was in NY, and the answer was that the
radioactivity present in dilute UAc (1 or 2%) was not high enough to
prohibit it being poured down the drain. YMMV, so check with your
safety office and anyone else who might have valuable input.
Yours,
Bill

==============================Original Headers==============================
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I am shocked! Uranyl is not only radioactive, it is a toxic and a heavy metal and as such accumulates in organisms.
It shouldn't be poured down the drain!
In my lab it is taken care of as a heavy metal contaminant (with lead and osmium) by a special company.

Regards,
Stephane
�
----- Original Message -----
X-from: "wtivol-at-sbcglobal.net" {wtivol-at-sbcglobal.net}
To: nizets2-at-yahoo.com
Cc:
Sent: Thursday, August 30, 2012 3:23 AM

On Aug 28, 2012, at 1:19 PM, William.H.Roberts-at-usa.dupont.com wrote:

} Dear Listserver,
}
} Is there anyone who can provide advice/instructions/warnings, etc.�
} with
} respect to proper (legal) disposal of uranyl acetate in the U.S.?
}
} Thanks, Bill Roberts

Dear Bill,
�� Since no one else has tackled this, I'll give it a shot.� My info is�
more than a decade old; however, at least this part is still valid:�
Different states have different rules, so the correct answer will�
depend on where you plan to dispose of the UAc.� When I was last�
confronted with this issue, I was in NY, and the answer was that the�
radioactivity present in dilute UAc (1 or 2%) was not high enough to�
prohibit it being poured down the drain.� YMMV, so check with your�
safety office and anyone else who might have valuable input.
�� Yours,
�� Bill

==============================Original Headers==============================
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17, 44 -- Subject: Re: [Microscopy] Re: Disposal of Uranyl Acetate - Help Needed
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Email: w.choi-at-bitplane.com
Name: Won Yung Choi

Organization: Bitplane

Title-Subject: [Filtered] Bitplane, a world leader in 3D/4D image analysis, is searching for a
talented regional sales engineer (RSE) to join a dynamic team of experts.

Message: Primary Responsibilities Include:

1. Meet/exceed agreed team and individual sales objectives for Bitplanes Imaris 3D/4D image
analysis software.
2. Establish new business relationships with key science centers and scientists, and strengthen
existing customer relationship within the defined territory (with a concentration in Houston, Texas).
3. Provide answers to technical and commercial queries from direct research customers.
4. Perform customized demonstration of Bitplane software in a manner tailored for customers
specific scientific endpoints to promote the sale of products.
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advanced optical microscopy/image analysis courses as commercial faculty.
6. Continually compile market information/intelligence on dedicated regions with a view to providing
market feedback to assist in the shaping of future company strategy and
7. Any other duties as may be reasonably required by the National Sales Manager.

Desired Qualifications:

1. Degree or equivalent in a science or engineering discipline (M.S. or Ph.D. is desired).
2. Expertise in confocal, widefield, live-cell imaging, and advanced image analysis.
3. Excellent communication skills.
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5. Creative problem solving ability.
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8. Experience in sales, application scientist or technical support role (in particular, in selling
or supporting technical high-value software).
9. Willingness to travel nationally and internationally as required.
10. Ability to speak Portuguese or Spanish (desired but not required).

If interested, please e-mail your resume and a cover letter to Rebecca Francoline
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Dr. Nestor J. Zaluzec
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Hello, Listserve members,

As with Bill's reply, I'm sure this may differ from state to state, and may differ from institution to institution. There is enough radon radiation in the water supply in Florida from all of the phosphate in our soil that our Health and Safety officers told us to flush our U.A., even our 8% stain solution, down the drain. We use such a small amount (0.5mL at a time, per stain run), that the dilution factor will render it harmless to the environment when compared to the amount of phosphate (and radon) present in river water. I'm sure this is correct. As an amateur paleontologist, I've put fossil ice-age mammal bone from my Florida adventures in my SEM before, turned on my EDS system with the E.M. beam off, and detected an x-ray signal, so I know the fossils are hot. I've been drinking Florida water for 32 years now..... No wonder I've been feeling kinda funny lately!

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046

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***************************************************************************************
Forwarded from "Ask a Microscopist"
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****************************************************************************************
} realname - Lauren Hale
} Email - lhale003-at-ucr,edu
} ORGANIZATION - University of California Riverside
} EDUCATION - Graduate College
} LOCATION - Riverside, CA, USA
} SUBJECT_OF_QUESTION - Dual confocal and SEM imagery
} QUESTION - Hello. Can you recommend some references or protocols
} for sample preparation to use a single sample for both confocal
} fluorescence microscopy and scanning electron microscopy? The
} sample will be a soil and root matrix and thus will need to be
} stabilized and used within 24 hours of prep (to visualize
} fluorescent signals). We have an environmental SEM so very little
} sample prep is needed for this, but the topology of the sample needs
} to be maintained. Any ideas or suggestions?

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We have had a similar experience with UA. Our radiation safety department
came and checked the radiation output of a bottle 2% uranyl acetate and
found the signal was barely above the background. It was so low that they
also said we did not have to collect the waste for disposal like we do
with most of our other waste from sample preparation and staining.

This may depend on where you live and the background presence of uranium.
I would also think it depended on the amount of waste produced. We
produce very little, maybe an ml a week of 2% UA when we are doing a lot
of negative staining. We usually use ~20�l droplets when staining grids.

Debby
---
Debby Sherman, Interim Director Phone: 765-494-6666
Life Science Microscopy Facility FAX: 765-494-5896

Purdue University E-mail: dsherman-at-purdue.edu

Debra Sherman, Chief Scientific Officer
DS imaging LLC
Purdue Technology Center
1281 Win Hentschel Blvd
West Lafayette, IN 47906
E-mail: debby.sherman-at-dsimagingllc.com
www.dsimagingllc.com
Mobile: 765-418-8540

On 8/30/12 8:25 AM, "ehaller-at-health.usf.edu" {ehaller-at-health.usf.edu}
wrote:

}
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Email: maryard-at-uga.edu
Name: Mary Ard

Organization: University of Georgia

Title-Subject: [Filtered] Magnifying Lens on Vibratome 1000 Plus

Message: Good Day. I am in need of replacing the magnifying lens on the Vibratome 1000 Plus we have
in our lab facility. I have not been able to locate the Vibratome Company - the manufacturer of the
Vibratome 1000 Plus - either by website or by phone. I am not sure they are currently in existence,
or if another company bought them out. I cannot justify the purchase of another vibratome just to
have the extra part, even though these models are out there in lab surplus supply areas. Is there
someone I may contact that would help locate this part? Thank you.

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Email: michael.cammer-at-med.nyu.edu
Name: Michael Cammer

Organization: NYU Langone Medical Center

Title-Subject: [Filtered] warning about fluorescent lens cleaner

Message: Just some minutiae to add to the lens cleaner lore. By accident (we were screening for
biological crud/spillage left by users) we found that Fisher brand lens cleaner is fluorescence.
Residue could lower the contrast of fluorescent imaging.
Picture at http://www.flickr.com/photos/mcammer/7894860042/
Regards,
Michael

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Email: ewood1-at-uvm.edu
Name: Erin Wood

Organization: University of Vermont-Mechanical Engineering

Title-Subject: [Filtered] JEOL JSM 6060 woes continued

Message: Hi everyone, I hope you are all in good health and spirits and I graciously thank all who
responded to my previous question regarding my jet assembly for the JEOL 6060 SEM. Sadly, our
microscope frustration continues....

we've reassembled the diffusion pump and it seems to have fit together properly. We added 100 mL of
Santovac-5 to the pump and reassembled it to the microscope. I ran the rough pump for a while and
then booted the SEM normally. It went through "Pre-evac" and started into "Evac" in the SEM
software but then it gave me an error saying "Code 167".

I do not know what this problem could be but it seems that it could be related to the operation of
valve v4. This is the gate valve. I have noticed that when I try to do an evacuation cycle this
valve stays open much longer than it has in our previous attempts to ressurect this microscope. I'm
sorry if this is a rather elementary question but I'm hoping that someone can explain to me what
would affect the performance of the gate valve especially why it wouldn't open or why it would stay
open.
Thank you in advance!

Erin L Wood

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Dear colleagues,
Many thanks to all those who spent some time to send me a reply.
I think I found a solution to my problem and I post it on the list for the archive, since it may help someone else someday.
Using a glass slide was not possible since I couldn't make them clean enough.
Fortunately I noticed that small plastic petri dishes used for cell culture are perfectly clean.
Unfortunately when I tried to dry 2�l of suspension on the petri dish, the liquid spread a lot (ethanol) and the surface to analyse was really too big.
Since the particles are sometimes quite small I require a minimum of 20x objective and I needed to stitch 540 fields together to get the whole picture :-D
I don't want to pipet less than 2�l because I think the pipetting error will increase too much.
So I am not able to quantify the whole volume (2�l).
Fortunately someone gave me the solution: just add a known concentration of latex beads to the powder suspension and you can relate the number of mineral particles to the number of latex beads (thus to the volume) anytime. I have the added advantage to be able to recognize specifically the latex beads through fluorescence, so it will make the task of separating them from the powder�very easy.
Once again the list has proved invaluable for me so thanks again.
�
Stephane

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Email: Angel.Paredes-at-uth.tmc.edu
Name: Angel Paredes

Organization: University of Texas Health Science Center

Title-Subject: [Filtered] Denton Desk V Evaporator won't go to high vac

Message: Hi,

I have a Denton Desk V carbon evaporator with a Turbo. The system was working fine but now does not
go to high vacuum. The operation appears to work fine without any notices of interlock failures or
any error messages. When controlled to go to high vac, it appears to work to get there but never
does. This is because the turbo never kicks on. Has anyone else experienced this problem? I
haven't checked any fuses yet but wanted to ask first if anyone else has had this problem with their
Denton Desktop V turbo and maybe has a better answer for this problem.

Thanks,
Angel Paredes

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Dear all,

� � I am looking for cheap and good quality formvar carbon film. I saw amazon has some formvar carbon films by GLOEMT. The price is very good. Has anyone ordered from this company before? It seems they have good amazon feedback and a few good customer product reviews. Below is a link for one of the products I am interested

� http://www.amazon.com/Formvar-stabilized-carbon-support-mounted/dp/B007HVWYBI/ref=sr_1_2?ie=UTF8&qid=1346719892&sr=8-2

� � Thank you!

Gary

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9, 37 -- From: Gary Zhang {zhanggary717-at-yahoo.com}
9, 37 -- Subject: Help with ordering formvar carbon film
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Email: hariteja2003-at-gmail.com
Name: Harikrishna

Organization: NCBS,Bangalore,INDIA

Title-Subject: [Filtered] Query regarding Fume hoods for TEM sampl preparation room

Message: Dear All,

I am Harikrishna writing form NCBS,Bangaore. We are setting up sample preparation room for TEM facility.

The query is: As of now we have thought of using HEPA filter and carbon filter before final exhaust
such that traces of heavy metal stain specifically Osmium Tetra oxide (if any) will be trapped in
carbon filter. The carbon filter after specified time will be incinerated. I wanted to know if this
method is appropriate or is there any better solution for this.

Could you please guide us on this.

Thanking you,

Kind Regards,
Harikrishna.
TEM facility,CIFF,
Bangalore,
INDIA.

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Dear Gary,

I have been ordering from them since March of this year. I have used their formvar stabilized with carbon support film A200. Films are clean and flat. No defect founded so far. Carbon film thickness is just right for me. It is thick enough to make films stable under the electron beam and thin enough to get good resolution images. By the way I work with biological samples.
I have also ordered the holey carbon films through their website. Works fine with me too. I�ve almost gone through the second box.
I will suggest you to start a small order first to see whether it works for you or not. I always do that when I order from a new company.
You can reach me by this email if you have additional question. I will be glad to share my experience with you. Hope this will help.

Maria

} Date: Mon, 3 Sep 2012 20:26:11 -0500
} To: Wbaby66-at-hotmail.com
} From: zhanggary717-at-yahoo.com
} Subject: [Microscopy] Help with ordering formvar carbon film
}
}
}
}
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} Dear all,
}
} I am looking for cheap and good quality formvar carbon film. I saw amazon has some formvar carbon films by GLOEMT. The price is very good. Has anyone ordered from this company before? It seems they have good amazon feedback and a few good customer product reviews. Below is a link for one of the products I am interested
}
} http://www.amazon.com/Formvar-stabilized-carbon-support-mounted/dp/B007HVWYBI/ref=sr_1_2?ie=UTF8&qid=1346719892&sr=8-2
}
} Thank you!
}
} Gary
}
}
}
}
}
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} 9, 37 -- Subject: Help with ordering formvar carbon film
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Organization: The New England Society for Microscopy

Title-Subject: [Filtered] Register Today: NESM Fall Meeting -at- UMass Amherst (Sept 28, 2012)

Message: Join NESM for our annual Fall Meeting on Friday, September 28 at UMass Amherst. The
meeting is composed of four concurrent afternoon workshops, a buffet dinner, and two technical talks.

Workshops:
-Introduction to HRSEM for Soft Materials
-Lessons Learned by Building a Multicolor TIRF/STORM Microscope
-Modern Raman Spectroscopy and Emerging Applications in Materials Science, Nano-Technology and Other
Fields
-Helium Ion Microscopy for Materials Application and Biological Imaging

Talks:
"Single-molecule-sensitive fluorescence microscopy of droplet-confined biomolecules", Lori Goldner,
Ph.D., UMass Amherst

Extraterrestrial metal as observed by electron microscopy techniques, Joseph Goldstein, Ph.D,
UMass Amherst

Register and find out more on our website: http://nesmicroscopy.org/upcoming-meeting.html

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Title-Subject: [Filtered] TEM Staining for Biological samples

Message: Dear Listener,
Some time negatively stained sample appears like positively stained. How this happens..??

What are the selection criteria for stains UA,PTA, Uranyl Format, Lead Citrate..??

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Hi Gary,

I have used their Formvar with carbon A200, Formvar with thick cabon B200 and lacey formvar with carbon E300 products. I am very happy with the products. Work well for me, no contamination, films are clean. I think it is worth a try.

Marry Gross

} Date: Mon, 3 Sep 2012 20:26:10 -0500
} To: reto12011-at-hotmail.com
} From: zhanggary717-at-yahoo.com
} Subject: [Microscopy] Help with ordering formvar carbon film
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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}
} Dear all,
}
} � � I am looking for cheap and good quality formvar carbon film. I saw amazon has some formvar carbon films by GLOEMT. The price is very good. Has anyone ordered from this company before? It seems they have good amazon feedback and a few good customer product reviews. Below is a link for one of the products I am interested
}
} � http://www.amazon.com/Formvar-stabilized-carbon-support-mounted/dp/B007HVWYBI/ref=sr_1_2?ie=UTF8&qid=1346719892&sr=8-2
}
} � � Thank you!
}
} Gary
}
}
}
}
}
} ==============================Original Headers==============================
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} 9, 37 -- From: Gary Zhang {zhanggary717-at-yahoo.com}
} 9, 37 -- Subject: Help with ordering formvar carbon film
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Hi Thomas,

Thank you very much for your input. I appreciate it. I am going to buy some to try.

Gary

--- On Mon, 9/3/12, THOMAS WILLIAMS {tomw-at-uidaho.edu} wrote:

} From: THOMAS WILLIAMS {tomw-at-uidaho.edu}
} Subject: Re: [Microscopy] Help with ordering formvar carbon film
} To: zhanggary717-at-yahoo.com
} Date: Monday, September 3, 2012, 6:59 PM
} Gary,
}
} Good evening.� I have used these carbon-stabilized
} grids for about a year now.� No problems and the
} quality is very good and very consistent.� I had my EM
} Class buy these and have heard no complaints so far.�
}
} I recommend that you give them a try.
}
} Cheers,
} Tom
}
} THOMAS WILLIAMS
} Asst. Dean & Director of Research Facilities
} College of Science
} University of Idaho
} Moscow, ID� 83844
} (208) 885-6785
} tomw-at-uidaho.edu
}
}
}
} On Sep 3, 2012, at 6:20 PM, zhanggary717-at-yahoo.com
} wrote:
}
} }
} }
} }
} }
} ----------------------------------------------------------------------------
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} }
} } Dear all,
} }
} } � ��I am looking for cheap and good
} quality formvar carbon film. I saw amazon has some formvar
} carbon films by GLOEMT. The price is very good. Has anyone
} ordered from this company before? It seems they have good
} amazon feedback and a few good customer product reviews.
} Below is a link for one of the products I am interested
} }
} } ��http://www.amazon.com/Formvar-stabilized-carbon-support-mounted/dp/B007HVWYBI/ref=sr_1_2?ie=UTF8&qid=1346719892&sr=8-2
} }
} } � ��Thank you!
} }
} } Gary
} }
} }
} }
} }
} }
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} } 9, 37 -- Subject: Help with ordering formvar carbon
} film
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9, 42 -- Subject: Re: [Microscopy] Help with ordering formvar carbon film
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Hello Ravi,
I think that the surface quality of supporting films is responsible for this. If you remove the stain around the particle by washing you get positively stained sample. However, when the supporting film is hydrophobic you can remove all of the stain by filter paper, too.

It can also happen when you negatively stain ribosomes or generally protein particles containing nucleic acid with uranyl acetate (uranyl formate) and remove almost all of the stain due to hydrophobic supporting film. Uranyl salts interact with nucleic acids and stain them positively.

Try to use glow-discharge activated carbon or carbon/formvar grids for negative staining. This can help you a lot.

Uranyl salt are generally used as unbuffered solutions (acidic). PTA solutions are usually titrated to neutral pH ~7.2 with KOH or NaOH.

Best regards from Prague Oldrich

--
Oldrich Benada
Institute of Microbiology, AS CR, v.v.i.
Videnska 1024
CZ-142 20 Prague 4
Czech Republic

On Wednesday 05 of September 2012 01:27:12 you wrote:
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} Title-Subject: [Filtered] TEM Staining for Biological samples
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} Some time negatively stained sample appears like positively stained. How this happens..??
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8, 40 -- From benada-at-biomed.cas.cz Wed Sep 5 02:26:51 2012
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Ravi,
If you can try to glow dishcharge your grids to make them more hydrophilic. Another trick is to use a wetting agent like 0.04% Tylose in your uranyl acetate. This will give you a more even staining.

Good Luck,
Michael Delannoy

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Email: amit.welcomes.u-at-gmail.com
Name: Amit Gupta

Organization: IISc

Title-Subject: [Filtered] correcting beam tilt(?)

Message: Good evening,
I work on JEM-2100f and have not-so-much experience on TEM. While aligning the beam I am bit
confused. If our gun tilt is ok and rotation center has been centred, then the caustic spot is not
symmetrical, rather it is like a circle with a bright dot on the side. On the other hand if I make
my caustic spot symmetrical using bright tilt than image shows some kind of band passing though
image near exact focus (see link) with severe astigmatism (on closer inspection it looks similar to
ring of infinite radial mag. in ronchigram). What exactly I am missing in alignment.
I exposed a formvar film for too long and saw that the burn mark also looks as if beam is tilted a
bit.
What I should focus more on? Rotation centring from HT wobbling or symmetric caustic spot? Should
they two not be the same?
Link to images: https://sites.google.com/site/auxilliarylinks/

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Hi

You clearly have a good question here as there does not seem to be a flood
of replies.

The first thing that I must say is it is impossible to have a variation in
astigmatism across an image the streaking that you see seems to be because
you are too near to condenser crossover when the illumination is incorrectly
aligned. I put myself in trouble with Nester if I ask you to look at my
Hints and Tips page but set out below is the Generic Alignment Data that I
believe is appropriate to you.

Try these alignment procedures because the problem has to be alignment.

Operating Instructions
for a JEOL 200C, 2000F, 2000FX

The { } indicates an optional/occasional alignment procedure depending on
the magnification you may be using?

1. Wait for the vacuum "ready" light
2. Check display and with HT OFF lower the accelerating voltage to
120kV Standing Current 120kV=60uA
3. Press HT
4. Very slowly, waiting each step for the current display to settle for
a couple of seconds, click to the desired kV � do not hold the switch in a
change position
5. Remove Specimen Holder and Objective aperture.
6. Increase the filament control to within half a division of the old
setting, if in doubt bring it up until there is a change in the emission
meter reading.
7. Reduce the magnification and adjust the condenser control to obtain
the smallest bright spot.
8. If the spot moves off centre re centre with ALIGNMENT SHIFT (on
desk)
9. Continue to adjust the condenser to form the smallest spot as you
increase the magnification until the image is magnified to be 2 to 3cms
across centre if required with ALIGNMENT SHIFT
10. Reduce the filament heating until the image breaks up.
11. Increase the filament heating to just obtain a spot and some halo.
12. Balance (have the intensity equal on opposite sides of the halo) the
spot and halo with the GUN TILT controls which are behind the right front
door, if the black area is equal all the way round this is fine.
13. Heat the filament to form more of a halo and re align GUN TILT to
balance this halo, repeat until you reach saturation. If the spot moves on
the screen use GUN SHIFT to re centre. When you saturate leave a few fine
marks in the spot.
14. With the condenser adjusted to form the smallest spot, saturate, and
centre the spot with the illumination ALIG SHIFT on the desk, spread the
condenser clockwise until the illumination is just short of filling the
screen.
15. Align the condenser aperture with its alignment controls to place
the beam exactly central on the screen.
16. Re check 14 and 15 for a constant centre
17. De saturate slightly, focus the filament image with the condenser
control and then with the condenser stigmator control X and Y.
18. Re saturate but leave a very slight dark mark on the image.
19. Check the emission current, how far is it above the Standing
Current? For tissue biology you need 10 to 20uA for virus work 20 to 30uA
for materials science 15 to 35uA.
20. To change the emission current turn the BIAS control which is on the
left hand desk
21. If you are lowering the current you will be able to turn the
filament down also, check the saturation with a spot and halo.
22. If you are raising the current check the spot and halo and re
saturate, re check the emission current.
23. Spread condenser CLOCKWISE. Set the filament stop. Turn the
filament down. Insert a specimen.
24. {Check for specimen movement when tilting and compensate.
25. Place a feature on the centre of the screen with the stage drives.
26. Release the tilt clamp, it pushes away. Tilt by 5 degs and re
centre the feature with the Z� control on the tilt mechanism
27. Tilt back to 0 and centre feature with the stage controls
28. Repeat for a constant centre LOCK THE UNIT AGAIN AT ZERO TILT}
29. Once you have carried out the above (24 to 28), once a week if you
do not intend to tilt the specimen, or if magnification accuracy is not
required -
30. EITHER
31. Switch on the wobbler X or Y when you insert a new specimen and
adjust the Z' knob on the tilt mechanism to bring the specimen into focus.
Otherwise for higher magnification accuracy you should check the
tilt/specimen movement each time you load a specimen.
32. OR
33. Set the Objective lens current to~7 amps at 200kV, switch on the
wobbler and adjust the Z� to obtain focus
34. Press LOW MAG, spread the illumination with the condenser coarse and
find a suitable area of the specimen.
35. {LM alignment - place a significant feature on the centre of the
screen in MAG but minimum setting. Press LOW MAG spread the condenser and
using the LOW MAG TILT (left front) bring the feature to the centre of the
screen. Correct the illumination centre with LOW MAG SHIFT.}
36. Press SA DIFF and trim to a bright spot with the Diff Focus control
and the condenser control
37. Insert and centre the objective aperture (specimen level) and centre
on the diffraction spot
38. Press MAG and adjust magnification, then set focus and illumination
and illumination centre to the desired levels
39. {TILT ALIGNMENT If the image moves at focus: press Objective lens
wobbler and adjust for minimum movement with the "bright tilt" controls, re
check 36. For work above 50,000X you need voltage alignment which again
requires the pulsing image to be centred on the screen with the �bright
tilt� controls}
40. Stigmator adjustment - (you need at least 50,000X to make a good
correction, go higher then come down to the mag you wish to use) focus for
maximum contrast and then use the objective stigmator X and Y as fine focus
controls, again aiming for maximum contrast amongst the fine structure.
41. To record an image on the TV system �
42. Lower the screen intensity and then raise the screen by the Screen L
button
43. Focus & Correct the astigmatism at a minimum of double the
magnification you wish for image recording, if there is a lens change STOP
and see the explanation below
44. If the image went through a lens change this means you are limited
to an increase in magnification no higher than this point to focus and
stigmate as a lens change may alter these conditions
45. Set the image recording magnification and proceed
46. Set the illumination to obtain the desired image brightness
47. Record the data
48. {Condenser Alignment Check (once each week) Check the "gun alignment
SHIFT" centre at spot size 1 with the condenser adjusted to form the
smallest spot
49. Check the "illumination alignment SHIFT" at spot size 3 with the
condenser adjusted to form the smallest spot
50. Repeat 47 and 48 for a constant centre}
51. {SPOT SIZE - for high performance work the spot should be about 2 to
5um in diameter check condenser cross over for size by going to 10,000X
where 1cm = 1um. You should use condenser aperture 2}

Day today Operation

1. Check the Basic Alignment as above
2. Lower the mag and spread the beam to fill the screen take out the
objective aperture� turn down the filament.
3. Insert a specimen
4. Check focus at ~7.03 amps and press wobbler X
5. Adjust the tilt Z� for focus, switch off wobbler
6. Go to DIFF and adjust Brightness and DIFF FOCUS to obtain a nice
spot
7. Insert and Centre the objective aperture
8. Go to higher magnification Focus & Stigmate the objective
9. Try to work with ~20uA emission
10. Above 50,000X check the HT wobbler centre. If it needed to be
changed check the centre of the diffraction spot and objective aperture
centre. Then check the illumination and condenser aperture centre.
11. When finished lower the mag and spread the beam to fill the screen �
turn down the filament
12. End of the day turn off the HT

Come back to me if you need more.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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Email: amit.welcomes.u-at-gmail.com
Name: Amit Gupta

Organization: IISc

Title-Subject: [Filtered] correcting beam tilt(?)

Message: Good evening,
I work on JEM-2100f and have not-so-much experience on TEM. While aligning
the beam I am bit confused. If our gun tilt is ok and rotation center has
been centred, then the caustic spot is not symmetrical, rather it is like a
circle with a bright dot on the side. On the other hand if I make my caustic
spot symmetrical using bright tilt than image shows some kind of band
passing though image near exact focus (see link) with severe astigmatism (on
closer inspection it looks similar to ring of infinite radial mag. in
ronchigram). What exactly I am missing in alignment.
I exposed a formvar film for too long and saw that the burn mark also
looks as if beam is tilted a bit.
What I should focus more on? Rotation centring from HT wobbling or symmetric
caustic spot? Should they two not be the same?
Link to images: https://sites.google.com/site/auxilliarylinks/

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Amit,

Just to point out that this is a FEGTEM - so the part in Steve's
protocol below about reducing the filament current doesn't apply, you
just use the anode wobbler instead and make the beam oscillation radial
about its' centre.

The 2100 can be difficult to align since it has a fairly sophisticated
condenser lens system - 3 condenser lenses plus a condenser mini-lens.
This means you can independently change the convergence angle (alpha
control) and beam dimensions (spot size). Although the alignment
procedure is nominally the same as the older two-lens machines, in
practice you may have to go round a loop a couple of times until
everything is aligned - i.e. when you align one parameter, another one
may become misaligned. It's more like 'tuning' than aligning.

The other very important thing is that you do the alignments at
'standard focus' (special button with a cover!!). Generally you don't
touch the focus apart from very fine tweaks, instead you set the machine
to standard focus and use the Z control to bring the specimen to the
right height (i.e. in focus). Also you need to be at 40,000x or above to
make sure all the projector lenses are switched on (some changes in
focus and alignment will be present at low mags, when some projector
lenses are turned off). Do this first and then follow the steps below.

1.) Because of the interaction between the different alignments, I
generally start with the Tilt centre. Choose your alpha (usually 3) and
spot size 1. Focus the beam to a spot, turn the Tilt-X wobbler on and
make the two spots one by selecting the Tilt align and using the Def-X
knob. If the spots move on parallel tracks when you do this and never
become one, select the Angle align to make the spots move in the
perpendicular direction, again with Def-X. You can centre the beam with
Shift-X while you're doing this, if you need to. Once you have a single
spot, turn Tilt X wobbler off, and repeat for Tilt Y.

2.) Check the Condenser aperture is centred and next try a voltage (HT)
centre using the HT wobbler and Bright Tilt -} Def-X,Y - if this is way
off, just make the beam oscillate concentrically as you did with the
anode wobbler. If it is not too far off, make the centre of image
stationary (tip - it's easier to find the mid-point of the /direction
/of the image movement, rather than the smallest amplitude of movement).
Once you've done this, check the condenser aperture again, and then the
anode wobbler again.

3.) Finally do the Gun shift alignment (the only alignment in which the
Shift-X and Y knobs change their function). Spot size 5 - centre with
Beam shift, then spot size 1, centre with Gun shift. Repeat until both
are centred. Go back to (2) and check they are still okay (will probably
have to do at least a HT centre).

Eventually you should get to the point where all the different
alignments are pretty close. There is one more alignment which can have
a big effect on everything else - the Shift centre - but hopefully this
is okay for you, if not make it #4 in the list above. If you change
alpha (or even go to Low Mag and back to Mag again) you may have to
repeat the alignment, usually only fairly small adjustments.

Finally at the end you can save the alignment in a file, so even if
someone else comes along and messes everything up, you can reload the
same lens currents and use it as a good starting point (but note this
will not work after a change of filament!!)

Hope this helps and good luck

Richard

-------- Original Message --------

Hi

You clearly have a good question here as there does not seem to be a flood
of replies.

The first thing that I must say is it is impossible to have a variation in
astigmatism across an image the streaking that you see seems to be because
you are too near to condenser crossover when the illumination is incorrectly
aligned. I put myself in trouble with Nester if I ask you to look at my
Hints and Tips page but set out below is the Generic Alignment Data that I
believe is appropriate to you.

Try these alignment procedures because the problem has to be alignment.

Operating Instructions
for a JEOL 200C, 2000F, 2000FX

The { } indicates an optional/occasional alignment procedure depending on
the magnification you may be using?

1. Wait for the vacuum "ready" light
2. Check display and with HT OFF lower the accelerating voltage to
120kV Standing Current 120kV=60uA
3. Press HT
4. Very slowly, waiting each step for the current display to settle for
a couple of seconds, click to the desired kV � do not hold the switch in a
change position
5. Remove Specimen Holder and Objective aperture.
6. Increase the filament control to within half a division of the old
setting, if in doubt bring it up until there is a change in the emission
meter reading.
7. Reduce the magnification and adjust the condenser control to obtain
the smallest bright spot.
8. If the spot moves off centre re centre with ALIGNMENT SHIFT (on
desk)
9. Continue to adjust the condenser to form the smallest spot as you
increase the magnification until the image is magnified to be 2 to 3cms
across centre if required with ALIGNMENT SHIFT
10. Reduce the filament heating until the image breaks up.
11. Increase the filament heating to just obtain a spot and some halo.
12. Balance (have the intensity equal on opposite sides of the halo) the
spot and halo with the GUN TILT controls which are behind the right front
door, if the black area is equal all the way round this is fine.
13. Heat the filament to form more of a halo and re align GUN TILT to
balance this halo, repeat until you reach saturation. If the spot moves on
the screen use GUN SHIFT to re centre. When you saturate leave a few fine
marks in the spot.
14. With the condenser adjusted to form the smallest spot, saturate, and
centre the spot with the illumination ALIG SHIFT on the desk, spread the
condenser clockwise until the illumination is just short of filling the
screen.
15. Align the condenser aperture with its alignment controls to place
the beam exactly central on the screen.
16. Re check 14 and 15 for a constant centre
17. De saturate slightly, focus the filament image with the condenser
control and then with the condenser stigmator control X and Y.
18. Re saturate but leave a very slight dark mark on the image.
19. Check the emission current, how far is it above the Standing
Current? For tissue biology you need 10 to 20uA for virus work 20 to 30uA
for materials science 15 to 35uA.
20. To change the emission current turn the BIAS control which is on the
left hand desk
21. If you are lowering the current you will be able to turn the
filament down also, check the saturation with a spot and halo.
22. If you are raising the current check the spot and halo and re
saturate, re check the emission current.
23. Spread condenser CLOCKWISE. Set the filament stop. Turn the
filament down. Insert a specimen.
24. {Check for specimen movement when tilting and compensate.
25. Place a feature on the centre of the screen with the stage drives.
26. Release the tilt clamp, it pushes away. Tilt by 5 degs and re
centre the feature with the Z� control on the tilt mechanism
27. Tilt back to 0 and centre feature with the stage controls
28. Repeat for a constant centre LOCK THE UNIT AGAIN AT ZERO TILT}
29. Once you have carried out the above (24 to 28), once a week if you
do not intend to tilt the specimen, or if magnification accuracy is not
required -
30. EITHER
31. Switch on the wobbler X or Y when you insert a new specimen and
adjust the Z' knob on the tilt mechanism to bring the specimen into focus.
Otherwise for higher magnification accuracy you should check the
tilt/specimen movement each time you load a specimen.
32. OR
33. Set the Objective lens current to~7 amps at 200kV, switch on the
wobbler and adjust the Z� to obtain focus
34. Press LOW MAG, spread the illumination with the condenser coarse and
find a suitable area of the specimen.
35. {LM alignment - place a significant feature on the centre of the
screen in MAG but minimum setting. Press LOW MAG spread the condenser and
using the LOW MAG TILT (left front) bring the feature to the centre of the
screen. Correct the illumination centre with LOW MAG SHIFT.}
36. Press SA DIFF and trim to a bright spot with the Diff Focus control
and the condenser control
37. Insert and centre the objective aperture (specimen level) and centre
on the diffraction spot
38. Press MAG and adjust magnification, then set focus and illumination
and illumination centre to the desired levels
39. {TILT ALIGNMENT If the image moves at focus: press Objective lens
wobbler and adjust for minimum movement with the "bright tilt" controls, re
check 36. For work above 50,000X you need voltage alignment which again
requires the pulsing image to be centred on the screen with the �bright
tilt� controls}
40. Stigmator adjustment - (you need at least 50,000X to make a good
correction, go higher then come down to the mag you wish to use) focus for
maximum contrast and then use the objective stigmator X and Y as fine focus
controls, again aiming for maximum contrast amongst the fine structure.
41. To record an image on the TV system �
42. Lower the screen intensity and then raise the screen by the Screen L
button
43. Focus & Correct the astigmatism at a minimum of double the
magnification you wish for image recording, if there is a lens change STOP
and see the explanation below
44. If the image went through a lens change this means you are limited
to an increase in magnification no higher than this point to focus and
stigmate as a lens change may alter these conditions
45. Set the image recording magnification and proceed
46. Set the illumination to obtain the desired image brightness
47. Record the data
48. {Condenser Alignment Check (once each week) Check the "gun alignment
SHIFT" centre at spot size 1 with the condenser adjusted to form the
smallest spot
49. Check the "illumination alignment SHIFT" at spot size 3 with the
condenser adjusted to form the smallest spot
50. Repeat 47 and 48 for a constant centre}
51. {SPOT SIZE - for high performance work the spot should be about 2 to
5um in diameter check condenser cross over for size by going to 10,000X
where 1cm = 1um. You should use condenser aperture 2}

Day today Operation

1. Check the Basic Alignment as above
2. Lower the mag and spread the beam to fill the screen take out the
objective aperture� turn down the filament.
3. Insert a specimen
4. Check focus at ~7.03 amps and press wobbler X
5. Adjust the tilt Z� for focus, switch off wobbler
6. Go to DIFF and adjust Brightness and DIFF FOCUS to obtain a nice
spot
7. Insert and Centre the objective aperture
8. Go to higher magnification Focus & Stigmate the objective
9. Try to work with ~20uA emission
10. Above 50,000X check the HT wobbler centre. If it needed to be
changed check the centre of the diffraction spot and objective aperture
centre. Then check the illumination and condenser aperture centre.
11. When finished lower the mag and spread the beam to fill the screen �
turn down the filament
12. End of the day turn off the HT

Come back to me if you need more.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

--
Richard Beanland

Director,
Integrity Scientific Ltd.

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Thanks Steve for replying, the streak i showed in image indeed show up
only when i increase the brightness. So it is from moving nearer to
condenser crossover.

Just for the academic curiosity, whats exactly happening?

and why the sample is blackening in uneven way? what i inferred from
your mail is that its because of gun tilt, is it?

On Fri, Sep 7, 2012 at 2:20 PM, {protrain-at-emcourses.com} wrote:
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi
}
} You clearly have a good question here as there does not seem to be a flood
} of replies.
}
} The first thing that I must say is it is impossible to have a variation in
} astigmatism across an image the streaking that you see seems to be because
} you are too near to condenser crossover when the illumination is incorrectly
} aligned. I put myself in trouble with Nester if I ask you to look at my
} Hints and Tips page but set out below is the Generic Alignment Data that I
} believe is appropriate to you.
}
} Try these alignment procedures because the problem has to be alignment.
}
} Operating Instructions
} for a JEOL 200C, 2000F, 2000FX
}
} The { } indicates an optional/occasional alignment procedure depending on
} the magnification you may be using?
}
} 1. Wait for the vacuum "ready" light
} 2. Check display and with HT OFF lower the accelerating voltage to
} 120kV Standing Current 120kV=60uA
} 3. Press HT
} 4. Very slowly, waiting each step for the current display to settle for
} a couple of seconds, click to the desired kV � do not hold the switch in a
} change position
} 5. Remove Specimen Holder and Objective aperture.
} 6. Increase the filament control to within half a division of the old
} setting, if in doubt bring it up until there is a change in the emission
} meter reading.
} 7. Reduce the magnification and adjust the condenser control to obtain
} the smallest bright spot.
} 8. If the spot moves off centre re centre with ALIGNMENT SHIFT (on
} desk)
} 9. Continue to adjust the condenser to form the smallest spot as you
} increase the magnification until the image is magnified to be 2 to 3cms
} across centre if required with ALIGNMENT SHIFT
} 10. Reduce the filament heating until the image breaks up.
} 11. Increase the filament heating to just obtain a spot and some halo.
} 12. Balance (have the intensity equal on opposite sides of the halo) the
} spot and halo with the GUN TILT controls which are behind the right front
} door, if the black area is equal all the way round this is fine.
} 13. Heat the filament to form more of a halo and re align GUN TILT to
} balance this halo, repeat until you reach saturation. If the spot moves on
} the screen use GUN SHIFT to re centre. When you saturate leave a few fine
} marks in the spot.
} 14. With the condenser adjusted to form the smallest spot, saturate, and
} centre the spot with the illumination ALIG SHIFT on the desk, spread the
} condenser clockwise until the illumination is just short of filling the
} screen.
} 15. Align the condenser aperture with its alignment controls to place
} the beam exactly central on the screen.
} 16. Re check 14 and 15 for a constant centre
} 17. De saturate slightly, focus the filament image with the condenser
} control and then with the condenser stigmator control X and Y.
} 18. Re saturate but leave a very slight dark mark on the image.
} 19. Check the emission current, how far is it above the Standing
} Current? For tissue biology you need 10 to 20uA for virus work 20 to 30uA
} for materials science 15 to 35uA.
} 20. To change the emission current turn the BIAS control which is on the
} left hand desk
} 21. If you are lowering the current you will be able to turn the
} filament down also, check the saturation with a spot and halo.
} 22. If you are raising the current check the spot and halo and re
} saturate, re check the emission current.
} 23. Spread condenser CLOCKWISE. Set the filament stop. Turn the
} filament down. Insert a specimen.
} 24. {Check for specimen movement when tilting and compensate.
} 25. Place a feature on the centre of the screen with the stage drives.
} 26. Release the tilt clamp, it pushes away. Tilt by 5 degs and re
} centre the feature with the Z� control on the tilt mechanism
} 27. Tilt back to 0 and centre feature with the stage controls
} 28. Repeat for a constant centre LOCK THE UNIT AGAIN AT ZERO TILT}
} 29. Once you have carried out the above (24 to 28), once a week if you
} do not intend to tilt the specimen, or if magnification accuracy is not
} required -
} 30. EITHER
} 31. Switch on the wobbler X or Y when you insert a new specimen and
} adjust the Z' knob on the tilt mechanism to bring the specimen into focus.
} Otherwise for higher magnification accuracy you should check the
} tilt/specimen movement each time you load a specimen.
} 32. OR
} 33. Set the Objective lens current to~7 amps at 200kV, switch on the
} wobbler and adjust the Z� to obtain focus
} 34. Press LOW MAG, spread the illumination with the condenser coarse and
} find a suitable area of the specimen.
} 35. {LM alignment - place a significant feature on the centre of the
} screen in MAG but minimum setting. Press LOW MAG spread the condenser and
} using the LOW MAG TILT (left front) bring the feature to the centre of the
} screen. Correct the illumination centre with LOW MAG SHIFT.}
} 36. Press SA DIFF and trim to a bright spot with the Diff Focus control
} and the condenser control
} 37. Insert and centre the objective aperture (specimen level) and centre
} on the diffraction spot
} 38. Press MAG and adjust magnification, then set focus and illumination
} and illumination centre to the desired levels
} 39. {TILT ALIGNMENT If the image moves at focus: press Objective lens
} wobbler and adjust for minimum movement with the "bright tilt" controls, re
} check 36. For work above 50,000X you need voltage alignment which again
} requires the pulsing image to be centred on the screen with the �bright
} tilt� controls}
} 40. Stigmator adjustment - (you need at least 50,000X to make a good
} correction, go higher then come down to the mag you wish to use) focus for
} maximum contrast and then use the objective stigmator X and Y as fine focus
} controls, again aiming for maximum contrast amongst the fine structure.
} 41. To record an image on the TV system �
} 42. Lower the screen intensity and then raise the screen by the Screen L
} button
} 43. Focus & Correct the astigmatism at a minimum of double the
} magnification you wish for image recording, if there is a lens change STOP
} and see the explanation below
} 44. If the image went through a lens change this means you are limited
} to an increase in magnification no higher than this point to focus and
} stigmate as a lens change may alter these conditions
} 45. Set the image recording magnification and proceed
} 46. Set the illumination to obtain the desired image brightness
} 47. Record the data
} 48. {Condenser Alignment Check (once each week) Check the "gun alignment
} SHIFT" centre at spot size 1 with the condenser adjusted to form the
} smallest spot
} 49. Check the "illumination alignment SHIFT" at spot size 3 with the
} condenser adjusted to form the smallest spot
} 50. Repeat 47 and 48 for a constant centre}
} 51. {SPOT SIZE - for high performance work the spot should be about 2 to
} 5um in diameter check condenser cross over for size by going to 10,000X
} where 1cm = 1um. You should use condenser aperture 2}
}
} Day today Operation
}
} 1. Check the Basic Alignment as above
} 2. Lower the mag and spread the beam to fill the screen take out the
} objective aperture� turn down the filament.
} 3. Insert a specimen
} 4. Check focus at ~7.03 amps and press wobbler X
} 5. Adjust the tilt Z� for focus, switch off wobbler
} 6. Go to DIFF and adjust Brightness and DIFF FOCUS to obtain a nice
} spot
} 7. Insert and Centre the objective aperture
} 8. Go to higher magnification Focus & Stigmate the objective
} 9. Try to work with ~20uA emission
} 10. Above 50,000X check the HT wobbler centre. If it needed to be
} changed check the centre of the diffraction spot and objective aperture
} centre. Then check the illumination and condenser aperture centre.
} 11. When finished lower the mag and spread the beam to fill the screen �
} turn down the filament
} 12. End of the day turn off the HT
}
} Come back to me if you need more.
}
} Steve
}
} Steve Chapman FRMS
} Protrain for Consultancy and Courses in Electron Microscopy
} Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
} www.emcourses.com
}
}
}
} -----Original Message-----
} X-from: microscopylistserver-noreply-at-microscopy.com
} [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: 06 September 2012 15:11
} To: protrain-at-emcourses.com
} Subject: [Microscopy] viaWWW: TEM correcting beam tilt(?)
}
}
}
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} Email: amit.welcomes.u-at-gmail.com
} Name: Amit Gupta
}
} Organization: IISc
}
} Title-Subject: [Filtered] correcting beam tilt(?)
}
} Message: Good evening,
} I work on JEM-2100f and have not-so-much experience on TEM. While aligning
} the beam I am bit confused. If our gun tilt is ok and rotation center has
} been centred, then the caustic spot is not symmetrical, rather it is like a
} circle with a bright dot on the side. On the other hand if I make my caustic
} spot symmetrical using bright tilt than image shows some kind of band
} passing though image near exact focus (see link) with severe astigmatism (on
} closer inspection it looks similar to ring of infinite radial mag. in
} ronchigram). What exactly I am missing in alignment.
} I exposed a formvar film for too long and saw that the burn mark also
} looks as if beam is tilted a bit.
} What I should focus more on? Rotation centring from HT wobbling or symmetric
} caustic spot? Should they two not be the same?
} Link to images: https://sites.google.com/site/auxilliarylinks/
}
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Hi

Yes the streaking on your image is due to the condenser lens being operated
too near to crossover. You should always work with the condenser set
clockwise from its focal point, under which condition the effect is that the
beam spot (the condenser cross over) becomes the virtual source. We are
trying to have the beam as parallel as possible so the smaller the spot size
and the further overfocus you run your system the better the quality of the
illumination and therefore the final image. If you are operating with too
little emission current the problem of working too near to crossover is very
common.

As has been pointed out I do not know your system but the alignment of the
condenser lenses to remove a dog leg from the beam path is always very
important. If this alignment is not achieved the misalignment of the gun
(moved off axis) is being corrected by the illumination system (moved back
on axis) rather than the beam travelling straight down the column though the
multiple condenser lenses. The off centre contamination demonstrates this
error.

I hope this helps further?

Regards

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: amit.welcomes.u-at-gmail.com [mailto:amit.welcomes.u-at-gmail.com]
Sent: 08 September 2012 05:59
To: protrain-at-emcourses.com

Thanks Steve for replying, the streak i showed in image indeed show up only
when i increase the brightness. So it is from moving nearer to condenser
crossover.

Just for the academic curiosity, whats exactly happening?

and why the sample is blackening in uneven way? what i inferred from your
mail is that its because of gun tilt, is it?

On Fri, Sep 7, 2012 at 2:20 PM, {protrain-at-emcourses.com} wrote:
}
}
}
} ----------------------------------------------------------------------
} ------ The Microscopy ListServer -- CoSponsor: The Microscopy Society
} of America To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------
} ------
}
} Hi
}
} You clearly have a good question here as there does not seem to be a
} flood of replies.
}
} The first thing that I must say is it is impossible to have a
} variation in astigmatism across an image the streaking that you see
} seems to be because you are too near to condenser crossover when the
} illumination is incorrectly aligned. I put myself in trouble with
} Nester if I ask you to look at my Hints and Tips page but set out
} below is the Generic Alignment Data that I believe is appropriate to you.
}
} Try these alignment procedures because the problem has to be alignment.
}
} Operating Instructions
} for a JEOL 200C, 2000F, 2000FX
}
} The { } indicates an optional/occasional alignment procedure
} depending on the magnification you may be using?
}
} 1. Wait for the vacuum "ready" light
} 2. Check display and with HT OFF lower the accelerating voltage to
} 120kV Standing Current 120kV=60uA
} 3. Press HT
} 4. Very slowly, waiting each step for the current display to settle
for
} a couple of seconds, click to the desired kV � do not hold the switch
} in a change position
} 5. Remove Specimen Holder and Objective aperture.
} 6. Increase the filament control to within half a division of the old
} setting, if in doubt bring it up until there is a change in the
} emission meter reading.
} 7. Reduce the magnification and adjust the condenser control to
obtain
} the smallest bright spot.
} 8. If the spot moves off centre re centre with ALIGNMENT SHIFT (on
} desk)
} 9. Continue to adjust the condenser to form the smallest spot as you
} increase the magnification until the image is magnified to be 2 to
} 3cms across centre if required with ALIGNMENT SHIFT
} 10. Reduce the filament heating until the image breaks up.
} 11. Increase the filament heating to just obtain a spot and some halo.
} 12. Balance (have the intensity equal on opposite sides of the halo)
the
} spot and halo with the GUN TILT controls which are behind the right
} front door, if the black area is equal all the way round this is fine.
} 13. Heat the filament to form more of a halo and re align GUN TILT to
} balance this halo, repeat until you reach saturation. If the spot moves
on
} the screen use GUN SHIFT to re centre. When you saturate leave a few
} fine marks in the spot.
} 14. With the condenser adjusted to form the smallest spot, saturate,
and
} centre the spot with the illumination ALIG SHIFT on the desk, spread
} the condenser clockwise until the illumination is just short of
} filling the screen.
} 15. Align the condenser aperture with its alignment controls to place
} the beam exactly central on the screen.
} 16. Re check 14 and 15 for a constant centre
} 17. De saturate slightly, focus the filament image with the condenser
} control and then with the condenser stigmator control X and Y.
} 18. Re saturate but leave a very slight dark mark on the image.
} 19. Check the emission current, how far is it above the Standing
} Current? For tissue biology you need 10 to 20uA for virus work 20 to
} 30uA for materials science 15 to 35uA.
} 20. To change the emission current turn the BIAS control which is on
the
} left hand desk
} 21. If you are lowering the current you will be able to turn the
} filament down also, check the saturation with a spot and halo.
} 22. If you are raising the current check the spot and halo and re
} saturate, re check the emission current.
} 23. Spread condenser CLOCKWISE. Set the filament stop. Turn the
} filament down. Insert a specimen.
} 24. {Check for specimen movement when tilting and compensate.
} 25. Place a feature on the centre of the screen with the stage drives.
} 26. Release the tilt clamp, it pushes away. Tilt by 5 degs and re
} centre the feature with the Z� control on the tilt mechanism
} 27. Tilt back to 0 and centre feature with the stage controls
} 28. Repeat for a constant centre LOCK THE UNIT AGAIN AT ZERO TILT}
} 29. Once you have carried out the above (24 to 28), once a week if you
} do not intend to tilt the specimen, or if magnification accuracy is
} not required -
} 30. EITHER
} 31. Switch on the wobbler X or Y when you insert a new specimen and
} adjust the Z' knob on the tilt mechanism to bring the specimen into focus.
} Otherwise for higher magnification accuracy you should check the
} tilt/specimen movement each time you load a specimen.
} 32. OR
} 33. Set the Objective lens current to~7 amps at 200kV, switch on the
} wobbler and adjust the Z� to obtain focus
} 34. Press LOW MAG, spread the illumination with the condenser coarse
and
} find a suitable area of the specimen.
} 35. {LM alignment - place a significant feature on the centre of the
} screen in MAG but minimum setting. Press LOW MAG spread the condenser
} and using the LOW MAG TILT (left front) bring the feature to the
} centre of the screen. Correct the illumination centre with LOW MAG
SHIFT.}
} 36. Press SA DIFF and trim to a bright spot with the Diff Focus
control
} and the condenser control
} 37. Insert and centre the objective aperture (specimen level) and
centre
} on the diffraction spot
} 38. Press MAG and adjust magnification, then set focus and
illumination
} and illumination centre to the desired levels
} 39. {TILT ALIGNMENT If the image moves at focus: press Objective lens
} wobbler and adjust for minimum movement with the "bright tilt"
} controls, re check 36. For work above 50,000X you need voltage
} alignment which again requires the pulsing image to be centred on the
} screen with the �bright tilt� controls}
} 40. Stigmator adjustment - (you need at least 50,000X to make a good
} correction, go higher then come down to the mag you wish to use) focus
} for maximum contrast and then use the objective stigmator X and Y as
} fine focus controls, again aiming for maximum contrast amongst the fine
structure.
} 41. To record an image on the TV system �
} 42. Lower the screen intensity and then raise the screen by the Screen
L
} button
} 43. Focus & Correct the astigmatism at a minimum of double the
} magnification you wish for image recording, if there is a lens change
} STOP and see the explanation below
} 44. If the image went through a lens change this means you are limited
} to an increase in magnification no higher than this point to focus and
} stigmate as a lens change may alter these conditions
} 45. Set the image recording magnification and proceed
} 46. Set the illumination to obtain the desired image brightness
} 47. Record the data
} 48. {Condenser Alignment Check (once each week) Check the "gun
alignment
} SHIFT" centre at spot size 1 with the condenser adjusted to form the
} smallest spot
} 49. Check the "illumination alignment SHIFT" at spot size 3 with the
} condenser adjusted to form the smallest spot
} 50. Repeat 47 and 48 for a constant centre}
} 51. {SPOT SIZE - for high performance work the spot should be about 2
to
} 5um in diameter check condenser cross over for size by going to
} 10,000X where 1cm = 1um. You should use condenser aperture 2}
}
} Day today Operation
}
} 1. Check the Basic Alignment as above
} 2. Lower the mag and spread the beam to fill the screen take out the
} objective aperture� turn down the filament.
} 3. Insert a specimen
} 4. Check focus at ~7.03 amps and press wobbler X
} 5. Adjust the tilt Z� for focus, switch off wobbler
} 6. Go to DIFF and adjust Brightness and DIFF FOCUS to obtain a nice
} spot
} 7. Insert and Centre the objective aperture
} 8. Go to higher magnification Focus & Stigmate the objective
} 9. Try to work with ~20uA emission
} 10. Above 50,000X check the HT wobbler centre. If it needed to be
} changed check the centre of the diffraction spot and objective
} aperture centre. Then check the illumination and condenser aperture
centre.
} 11. When finished lower the mag and spread the beam to fill the screen
�
} turn down the filament
} 12. End of the day turn off the HT
}
} Come back to me if you need more.
}
} Steve
}
} Steve Chapman FRMS
} Protrain for Consultancy and Courses in Electron Microscopy Tel & Fax
} +44 (0)1280 816512 Cell +44 (0)7711 606967 www.emcourses.com
}
}
}
} -----Original Message-----
} X-from: microscopylistserver-noreply-at-microscopy.com
} [mailto:microscopylistserver-noreply-at-microscopy.com]
} Sent: 06 September 2012 15:11
} To: protrain-at-emcourses.com
} Subject: [Microscopy] viaWWW: TEM correcting beam tilt(?)
}
}
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} Email: amit.welcomes.u-at-gmail.com
} Name: Amit Gupta
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} Organization: IISc
}
} Title-Subject: [Filtered] correcting beam tilt(?)
}
} Message: Good evening,
} I work on JEM-2100f and have not-so-much experience on TEM. While
} aligning the beam I am bit confused. If our gun tilt is ok and
} rotation center has been centred, then the caustic spot is not
} symmetrical, rather it is like a circle with a bright dot on the side.
} On the other hand if I make my caustic spot symmetrical using bright tilt
than image shows some kind of band
} passing though image near exact focus (see link) with severe
} astigmatism (on closer inspection it looks similar to ring of infinite
} radial mag. in ronchigram). What exactly I am missing in alignment.
} I exposed a formvar film for too long and saw that the burn mark
} also looks as if beam is tilted a bit.
} What I should focus more on? Rotation centring from HT wobbling or
} symmetric caustic spot? Should they two not be the same?
} Link to images: https://sites.google.com/site/auxilliarylinks/
}
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Hi David

Someone else might have a more specific answer to your question but this might get you started:
First try the vendors of the equipment to see if they provide general training.
Also, we have used input from biomedical illustrators, they are generally photographers who have specialised.

http://www.aimbi.org.au/links/ This is an Aus website that provides some international links.

https://www.lifesci.ucsb.edu/mcdb/events/imaging_workshop/ This workshop looked good but distance prevented us attending. It is run annually.

Hope this is of some help,
Naomi

} } } {oshel1pe-at-cmich.edu} 9/7/2012 10:52 pm } } }

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} realname - David O'Dowd
} Email - david.odowd-at-stemcell.com
} LOCATION - Vancouver BC
} SUBJECT_OF_QUESTION - Microscopy Eduation
} QUESTION - Hi there,
} My name is David,
} I head up the video dept for STEMCELL Technologies here in Vancouver
} BC Canada. We are a small group within the company that is
} responsible for capturing creating all video assets in our company.
} The problem being that in the dept, we are all Video experts, not
} scientists, this creates a problem when it comes to our
} understanding of Photomicrography. It's difficult to find
} information on the topic that speaks to photographers rather than
} Scientists. Would anybody in your organization know of any resources
} or education programs that speak to our particular issue?
}
} Thanks in advance for any reply.
} -David O'Dowd

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16, 28 -- From prvs=1600037094=Naomi_McCallum-at-health.qld.gov.au Sun Sep 9 22:54:58 2012
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Someone from another department came by today, asking if I could look at a number of images of blood clots in blood vessels taken using SEM. They were wondering if I could differentiate platelets from white cells. While I could probably guess by size, I am for most part out of my experience here. Is there someone (preferably with experience in interpreting these types of images) on the list that might be willing to work with this lab and help identify what is going on?

Send me a short email and I will pass your information along.

Suggested references might be nice as well. Thanks!

Doug

^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^
Douglas W. Cromey, M.S. - Associate Scientific Investigator
Dept. of Cellular & Molecular Medicine, University of Arizona
1501 N. Campbell Ave, Tucson, AZ 85724-5044 USA

office: AHSC 4212 email: Cromey-at-Arizona.edu
voice: 520-626-2824 fax: 520-626-2097

http://swehsc.pharmacy.arizona.edu/exppath/
Home of: "Microscopy and Imaging Resources on the WWW"

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10, 31 -- Subject: Interpreting SEM images of blood cells and clots
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Greeting

Over the week end we defrosted the refrigerator that stores our OsO4. Wondering if you have any advice regarding removing the typical black stains on the refrigerator walls. Ours are not bad, we triple contain our solutions, just curious if anyone has tried removing the stains or if it is better to 'let sleeping dogs lie'.

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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Hi Doug

Here's a start http://www.reading.ac.uk/CFAM/Imageofthemonth/Imageofthemonth2010/CfAM_June_2010.aspx
Try contacting this group. Unfortunately my experience only extends to diagnostic applications of TEM with platelets/WBC and a little RBC SEM.

Cheers
Naomi
} } } {dcromey-at-email.arizona.edu} 9/11/2012 3:50 am } } }

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Someone from another department came by today, asking if I could look at a number of images of blood clots in blood vessels taken using SEM. They were wondering if I could differentiate platelets from white cells. While I could probably guess by size, I am for most part out of my experience here. Is there someone (preferably with experience in interpreting these types of images) on the list that might be willing to work with this lab and help identify what is going on?

Send me a short email and I will pass your information along.

Suggested references might be nice as well. Thanks!

Doug

^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^
Douglas W. Cromey, M.S. - Associate Scientific Investigator
Dept. of Cellular & Molecular Medicine, University of Arizona
1501 N. Campbell Ave, Tucson, AZ 85724-5044 USA

office: AHSC 4212 email: Cromey-at-Arizona.edu
voice: 520-626-2824 fax: 520-626-2097

http://swehsc.pharmacy.arizona.edu/exppath/
Home of: "Microscopy and Imaging Resources on the WWW"

==============================Original Headers==============================
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PS Fibrin fibers are brown; platelet aggregates are gray; red blood cells are red; leukocytes are green.
http://news.pennmedicine.org/blog/2012/07/taking-advantage-of-mother-nature-delivering-drugs-using-red-blood-cells-.html

Sorry folks, just couldn't contain myself. Everything is coloured these days!

Cheers
Naomi
} } } {dcromey-at-email.arizona.edu} 9/11/2012 3:50 am } } }

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Someone from another department came by today, asking if I could look at a number of images of blood clots in blood vessels taken using SEM. They were wondering if I could differentiate platelets from white cells. While I could probably guess by size, I am for most part out of my experience here. Is there someone (preferably with experience in interpreting these types of images) on the list that might be willing to work with this lab and help identify what is going on?

Send me a short email and I will pass your information along.

Suggested references might be nice as well. Thanks!

Doug

^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^
Douglas W. Cromey, M.S. - Associate Scientific Investigator
Dept. of Cellular & Molecular Medicine, University of Arizona
1501 N. Campbell Ave, Tucson, AZ 85724-5044 USA

office: AHSC 4212 email: Cromey-at-Arizona.edu
voice: 520-626-2824 fax: 520-626-2097

http://swehsc.pharmacy.arizona.edu/exppath/
Home of: "Microscopy and Imaging Resources on the WWW"

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19, 28 -- From prvs=1600037094=Naomi_McCallum-at-health.qld.gov.au Mon Sep 10 18:32:08 2012
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Just wanted to bring to the attention of anyone interested in a possible post-doctoral position at Pacific Northwest National Laboratory in Washington State.

Job ID: 301812
Directorate: Energy and Environment
Division: Radiological & Nuclear S & T
Group: Materials & Structures Performance

You can view and apply for this job at:
https://erecruit.pnnl.gov/psp/hrext/EMPLOYEE/HRMS/c/HRS_HRAM.HRS_CE.GBL?Page=HRS_CE_JOB_DTL&Action=A&JobOpeningId=301812&SiteId=1&PostingSeq=1

Job Description

The Engineering Mechanics and Structural Materials Group is seeking a Materials Science postdoctoral appointee to work on characterization of nanoscale features in structural materials. Prior experience in neutron-irradiated structural materials such as martensitic/ferritic steels, ODS ferritic/martensitic steels, and gas bubble/void formation is preferred but not essential in considerations for this position. The main duties of the successful candidate will involve the operation of a new aberration-corrected TEM/STEM devoted to characterizing irradiated materials, and prior experience with such instrumentation will be strong factor in being considered for the position. Primary job activities will include high-resolution characterization of neutron-irradiated metallic alloys and analysis of complex structures created during thermomechanical treatments and/or irradiation. The candidate must be capable of correlating physical and mechanical properties to the measured microstructural characteristics. Initial emphasis for the applicant involves working closely with scientists studying radiation effects in various structural materials for advanced nuclear energy systems. The candidate must be willing to travel and interact with scientists from across the world, and will be expected to publish their results in appropriate peer-reviewed journals and attend relevant conferences. The group's research is interdisciplinary in nature, including work related to radiation damage, high-temperature degradation, stress corrosion cracking, materials for transportation needs, solid oxide fuel cell development, hydrogen compatibility, and radiation detection.

Thank you,

Dan Edwards
Scientist
Engineering Mechanics and Structural Materials
Pacific Northwest National Laboratory
902 Battelle Boulevard
P.O. Box 999, MSIN J4-55
Richland, WA� 99352 USA
Tel:� 509-371-7284
Fax: 509-375-3033
dan.edwards-at-pnnl.gov
www.pnnl.gov

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Dear Doug,
Bear in mind that ideally you'd know if the platelets are activated or not...they go from roundish (a bit like erythrocytes=2C but smaller�to spiky with long pseudopods when fully activated.They are quite different to white cells which are a more spherical shape in normal human blood.
Regards,
Pete

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Title-Subject: [Filtered] TEM - Sorvall MT6000

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Email: harvey.guthrey-at-nrel.gov
Name: Harvey Guthrey

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Title-Subject: [Filtered] Heated/Cooled SEM Stage

Message: I need a SEM stage capable of being cooled to -196C and heated
to ~400C. The heating is less important than the cooling but would be
nice to have. Does anyone have any suggestions on which companies to
contact regarding this, or have a surplus stage with these capabilities?

Thanks

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Hi folks,

I have a client who would like to coat a silicon wafer with 10 microns
of Mg metal. I cannot do this with our system but was wondering if
anyone knows how we could this around the Minneapolis area?

Nick

--

Dr Nicholas Seaton

SEM Specialist
Characterization Facility

College of Science and Engineering

University of Minnesota

Shepherd Labs

100 Union Street SE

Minneapolis

MN, 55455

email: seato008-at-umn.edu

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Email: kevin-at-semionco.com
Name: Kevin Filter

Organization: Semion LLC

Title-Subject: [Filtered] schematics for Jeol JSM T200

Message: Hello, I recently acquired Jeol JSM T200 and it is in good condition except that the scans
are distorted. I do not have any schematics. Is there someone with schematics for this instrument
who is willing to share them? Maybe we can make a trade for services or some other item. Thanks for
your consideration. Kevin
semionco.com

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Email: tprozoro-at-ameslab.gov
Name: Tanya Prozorov

Organization: Ames Laboratory

Title-Subject: [Filtered] [STEM] QF grids problem

Message: Dear Colleagues,

Years ago I switched to the Quantfoil grids for the analysis of on-the-grid synthesized
nanoparticles, largely because of quality and stability of these grids.

The last order of these grids was a sore disappointment. I ordered several boxes of these from 2SPI,
and all but two boxes show heavy surface contamination. Their surface shows a pattern formed by a
residue of some sort, which interferes with my analysis. I contacted 2SPI folks, who, in turn are
in contact with QF people. Some grids are gold, some copper, with different mesh size, array of
holes and the Lot Numbers, so tracing the problem to a single bad batch might be a challenge.

I was wondering if anybody encountered similar problem. Meanwhile I am trying to figure out what the
residue is, and whether it is possible to remove it without causing too much damage to the carbon
film. I acquired quite a few images of the empty grids showing very annoying pattern, if anyone is
interested.

Any suggestion would be greatly appreciated.

Thank you,

-Tanya

Tanya Prozorov, Ph.D.

Emergent Atomic and Magnetic Structures
Division of Materials Sciences and Engineering, Ames Laboratory
136 B Wilhelm Hall, Ames IA 50011
515-2943376 (office)
515-2941255 (lab)
515-5207456 (cell)
515-294-8727 (fax)
e-mail: tprozoro-at-ameslab.gov
CREATING MATERIALS AND ENERGY SOLUTIONS

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PostDoc in cryo-electron tomography at C-CINA (University Basel) Basel, Switzerland.

A post-doctoral position to study the molecular mechanisms of the development of Parkinson's disease by cryo-electron microscopy is available in the laboratory of Henning Stahlberg, C-CINA, Biozentrum, University of Basel, Switzerland (http://c-cina.org) in a collaboration with F. Hoffmann La-Roche in Basel, and the group of Prof. Markus Tolnay, Pathology, University Basel Hospital. The position is funded initially for two years by the Roche Research Fellowship Program.

A PhD in Biology, Physics, or a related discipline, and experience in cryo-electron microscopy is required. Experience with high-pressure freezing, cryo-sectioning, electron tomography and/or image analysis is welcome.

The C-CINA laboratory enjoys outstanding equipment and has a pleasant working atmosphere. The lab language is English. Basel is a vibrant, cultural city with an excellent living quality at the Swiss border towards France and Germany. This post-doctoral position is at the interface between structural biology, medicine, and research in the pharmaceutical industry, and will give the successful candidate access to expertise, resources, and cultural atmosphere of these three institutions.

The position is available immediately. To apply, or for more information, please contact Henning Stahlberg, +41 61 387 32 62, Henning.Stahlberg-at-unibas.ch.

Henning Stahlberg, PhD
Prof. for Structural Biology, C-CINA, Biozentrum, University Basel
Mattenstrasse 26 | D-BSSE | WRO-1058 | CH-4058 Basel | Switzerland
http://c-cina.org | Tel. +41-61-387 32 62 | mailto:Henning.Stahlberg-at-unibas.ch

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On Sep 11, 2012, at 5:11 PM, microscopylistserver-
noreply-at-microscopy.com wrote:

} Name: Tanya Prozorov
}
} Organization: Ames Laboratory
}
} Title-Subject: [Filtered] [STEM] QF grids problem
}
} Message: Dear Colleagues,
}
} Years ago I switched to the Quantfoil grids for the analysis of on-
} the-grid synthesized
} nanoparticles, largely because of quality and stability of these
} grids.
}
} The last order of these grids was a sore disappointment. I ordered
} several boxes of these from 2SPI,
} and all but two boxes show heavy surface contamination. Their
} surface shows a pattern formed by a
} residue of some sort, which interferes with my analysis. I
} contacted 2SPI folks, who, in turn are
} in contact with QF people. Some grids are gold, some copper, with
} different mesh size, array of
} holes and the Lot Numbers, so tracing the problem to a single bad
} batch might be a challenge.
}
} I was wondering if anybody encountered similar problem. Meanwhile I
} am trying to figure out what the
} residue is, and whether it is possible to remove it without causing
} too much damage to the carbon
} film. I acquired quite a few images of the empty grids showing very
} annoying pattern, if anyone is
} interested.
}
} Any suggestion would be greatly appreciated.
}
} Thank you,
}
} -Tanya
}
} Tanya Prozorov, Ph.D.
}
} Emergent Atomic and Magnetic Structures
} Division of Materials Sciences and Engineering, Ames Laboratory
} 136 B Wilhelm Hall, Ames IA 50011
} 515-2943376 (office)
} 515-2941255 (lab)
} 515-5207456 (cell)
} 515-294-8727 (fax)
} e-mail: tprozoro-at-ameslab.gov
} CREATING MATERIALS AND ENERGY SOLUTIONS
}
Dear Tanya,
Some years ago I had the same experience with Quantifoils. I think
that the residue is from incomplete removal of the formvar from the
carbon film. In order to remove this residue, you must be careful not
to touch the grid with the solvent, which I remember to be benzene,
but maybe chloroform or methylene chloride would be better. Put a
disk of 3MM filter paper into the bottom of a glass petri dish ~10 cm
in diameter, set the dish in a hood with one edge on top of something
about 1 cm high--many hoods have a step near the window, and the petri
dish can have one side on top of the step with the other edge on the
floor of the hood. Place grid(s) on the filter paper near the highest
point, pour a small amount of solvent near the lowest point, and cover
the dish. Leave overnight. The idea is to let the solvent be drawn
up the filter paper to the grid(s), where it will dissolve the residue
and deposit it into the paper.
Yours,
Bill

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Dear Bill:

Thank you for the suggestion, we are going to try this as soon as the schedule permits, probably with methylene chloride.

-Tanya

---------------------------------------h-----------------------------------------
Tanya Prozorov, Ph.D.

Emergent Atomic and Magnetic Structures
Division of Materials Sciences and Engineering, Ames Laboratory
136 B Wilhelm Hall, Ames IA 50011
515-2943376 (office)
515-2941255 (lab)
515-5207456 (cell)
515-294-8727 (fax)
e-mail: tprozoro-at-ameslab.gov
CREATING MATERIALS AND ENERGY SOLUTIONS

********THIS MESSAGE IS A PRIVATE COMMUNICATION AND AS SUCH
CONTENTS ARE EXEMPT FROM THE IOWA OPEN RECORDS ACT********

-----Original Message-----
X-from: Bill & Sue Tivol [mailto:wtivol-at-sbcglobal.net]
Sent: Wednesday, September 12, 2012 8:56 PM
To: microscopy-at-microscopy.com; tprozoro-at-ameslab.gov

Hi Tanya,

I believe that the common solvents for formvar are chloroform and
dichloroethane. I use the latter to dissolve the formvar powder when I
cast holey formvar/carbon films.

Not being an organic chemist, I can't say how well the methylene
chloride will work.

Good luck!
Henk

At 9/12/2012 10:01 PM, tprozoro-at-ameslab.gov wrote:
}
}
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} Dear Bill:
}
} Thank you for the suggestion, we are going to try this as soon as the schedule permits, probably with methylene chloride.
}
} -Tanya
}
} ---------------------------------------h-----------------------------------------
} Tanya Prozorov, Ph.D.
}
} Emergent Atomic and Magnetic Structures
} Division of Materials Sciences and Engineering, Ames Laboratory
} 136 B Wilhelm Hall, Ames IA 50011
} 515-2943376 (office)
} 515-2941255 (lab)
} 515-5207456 (cell)
} 515-294-8727 (fax)
} e-mail: tprozoro-at-ameslab.gov
} CREATING MATERIALS AND ENERGY SOLUTIONS
}
} ********THIS MESSAGE IS A PRIVATE COMMUNICATION AND AS SUCH
} CONTENTS ARE EXEMPT FROM THE IOWA OPEN RECORDS ACT********
}
}
} -----Original Message-----
} X-from: Bill & Sue Tivol [mailto:wtivol-at-sbcglobal.net]
} Sent: Wednesday, September 12, 2012 8:56 PM
} To: microscopy-at-microscopy.com; tprozoro-at-ameslab.gov
} Subject: Re: [Microscopy] viaWWW:[STEM] QF grids problem
}
}
} On Sep 11, 2012, at 5:11 PM, microscopylistserver- noreply-at-microscopy.com wrote:
}
} } Name: Tanya Prozorov
} }
} } Organization: Ames Laboratory
} }
} } Title-Subject: [Filtered] [STEM] QF grids problem
} }
} } Message: Dear Colleagues,
} }
} } Years ago I switched to the Quantfoil grids for the analysis of on-
} } the-grid synthesized nanoparticles, largely because of quality and
} } stability of these grids.
} }
} } The last order of these grids was a sore disappointment. I ordered
} } several boxes of these from 2SPI, and all but two boxes show heavy
} } surface contamination. Their surface shows a pattern formed by a
} } residue of some sort, which interferes with my analysis. I contacted
} } 2SPI folks, who, in turn are in contact with QF people. Some grids are
} } gold, some copper, with different mesh size, array of holes and the
} } Lot Numbers, so tracing the problem to a single bad batch might be a
} } challenge.
} }
} } I was wondering if anybody encountered similar problem. Meanwhile I am
} } trying to figure out what the residue is, and whether it is possible
} } to remove it without causing too much damage to the carbon film. I
} } acquired quite a few images of the empty grids showing very annoying
} } pattern, if anyone is interested.
} }
} } Any suggestion would be greatly appreciated.
} }
} } Thank you,
} }
} } -Tanya
} }
} } Tanya Prozorov, Ph.D.
} }
} } Emergent Atomic and Magnetic Structures Division of Materials Sciences
} } and Engineering, Ames Laboratory
} } 136 B Wilhelm Hall, Ames IA 50011
} } 515-2943376 (office)
} } 515-2941255 (lab)
} } 515-5207456 (cell)
} } 515-294-8727 (fax)
} } e-mail: tprozoro-at-ameslab.gov
} } CREATING MATERIALS AND ENERGY SOLUTIONS
} }
} Dear Tanya,
} Some years ago I had the same experience with Quantifoils. I think that the residue is from incomplete removal of the formvar from the carbon film. In order to remove this residue, you must be careful not to touch the grid with the solvent, which I remember to be benzene, but maybe chloroform or methylene chloride would be better. Put a disk of 3MM filter paper into the bottom of a glass petri dish ~10 cm in diameter, set the dish in a hood with one edge on top of something about 1 cm high--many hoods have a step near the window, and the petri dish can have one side on top of the step with the other edge on the floor of the hood. Place grid(s) on the filter paper near the highest point, pour a small amount of solvent near the lowest point, and cover the dish. Leave overnight. The idea is to let the solvent be drawn up the filter paper to the grid(s), where it will dissolve the residue and deposit it into the paper.
} Yours,
} Bill
}
}
}
}
}
}
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} 15, 32 -- From tprozoro-at-ameslab.gov Wed Sep 12 21:01:15 2012
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

==============================Original Headers==============================
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Hi All,

For the last couple of months I have being trying to label bacterial cell
walls (presumably outermost layer) with electron dense 0.8 nm diameter
clusters that should be covalently attached. I started imaging
cluster-labeled bacteria using 400 mesh formvar- and carbon-coated Cu
grids using a Tecnai 12 at 120kV. We were not able to see any labeling
particles.

Sample prep: Placing sample on glow-discharged grid, let it dry for a few
minutes, and
wick off excess solution.

We carried on the recommendation of imaging our samples using 5 nm Si grid
(9 windows) with the same scope. These grids do not have a great affinity
for whole-cell bacteria, any recommendations? Also, at high magnifications
(above 100 k) it is very difficult distinguish bacteria (lightly stained
with uranyl acetate) from the grid material. Any suggestions on how to
stain bacteria in order to avoid interference with the nanocluster
labeling?

I would greatly appreciate any guidance or suggestions.

Best wishes,
Frances

Frances P. Rodriguez-Rivera
Department of Chemistry
UC Berkeley

==============================Original Headers==============================
7, 24 -- From frodriguez-at-berkeley.edu Thu Sep 13 16:22:42 2012
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7, 24 -- To: {Microscopy-at-Microscopy.Com}
7, 24 -- Subject: TEM - Staining Cells, Nanocluster labeling
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Hi Frances,

Bacteria are easy to image without stain. If you want to see nanoparticles clusters on bacteria do not use stain it will make visibility very difficult to decide what is stain and what is label--sometimes hard without stain in some anaerobes.

I would start with cells on glow discharged continuous carbon films. Most bacteria are easy to image at 5-15 kx, Are you looking at special nanobacteria that you are using such high mags? What size are your nanoparticle labels? Can you image them on carbon films without bacteria--always a good sanity check.

I am only a few buildings away if you want to chat.

Roseann Csencsits, PhD
Scientist in Charge, Downing TEM Facility
Lawrence Berkeley Lab 01-365
1 Cyclotron Road
Berkeley, CA 94720
510-486-4548

On Sep 13, 2012, at 2:31 PM, frodriguez-at-berkeley.edu wrote:

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} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi All,
}
} For the last couple of months I have being trying to label bacterial cell
} walls (presumably outermost layer) with electron dense 0.8 nm diameter
} clusters that should be covalently attached. I started imaging
} cluster-labeled bacteria using 400 mesh formvar- and carbon-coated Cu
} grids using a Tecnai 12 at 120kV. We were not able to see any labeling
} particles.
}
} Sample prep: Placing sample on glow-discharged grid, let it dry for a few
} minutes, and
} wick off excess solution.
}
} We carried on the recommendation of imaging our samples using 5 nm Si grid
} (9 windows) with the same scope. These grids do not have a great affinity
} for whole-cell bacteria, any recommendations? Also, at high magnifications
} (above 100 k) it is very difficult distinguish bacteria (lightly stained
} with uranyl acetate) from the grid material. Any suggestions on how to
} stain bacteria in order to avoid interference with the nanocluster
} labeling?
}
} I would greatly appreciate any guidance or suggestions.
}
} Best wishes,
} Frances
}
} Frances P. Rodriguez-Rivera
} Department of Chemistry
} UC Berkeley

==============================Original Headers==============================
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Greetings,
I am building a freeze dryer for samples frozen in
tert-butanol (for SEM -- alternative to critcal point drying). I am
having difficulty finding information about the resistance of typical
vacuum pump oils to tert butanol. The pump will be a common rotary
vane pump and there will be foreline trap. Still, it seems reasonable
to use an oil that shows good resistance to tBA. Any leads in finding
one will be gratefully appreciated.

Many thanks!
Tobias
--
_ ____ __ ____
/ \ / / \ / \ \ Tobias I. Baskin
/ / / / \ \ \ Professor
/_ / __ /__ \ \ \__ Biology Department
/ / / \ \ \ 611 N. Pleasant St.
/ / / \ \ \ University of
Massachusetts
/ / ___ / \ \__/ \ ____ Amherst, MA, 01003
www.bio.umass.edu/biology/baskin
Voice: 413 - 545 - 1533 Fax: 413 - 545 - 3243

==============================Original Headers==============================
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Hi Tobias,

There are oils fluorinated oils (Fomblin being the prime example)
that were made to be resistant to corrosive gases and the like pumped
through standard rotary vane pumps. However, I think the more important
factor would be specifying a pump equipped with a gas ballast for high
vapour throughput. The tert-Butanol will likely mix with the pump oil,
but the relatively high oil temperature and sufficient ballasting should
allow the pump to clear out anything that makes it past the trap.

Some manufacturers make pumps that are designed with high
condensible load in mind. Edwards RV series pumps have a high
throughput gas ballast, and Welch makes a pump model specifically for
pumping in freeze dryer applications. They may be worth looking into.

-Jacob

On 12-09-13 03:34 PM, baskin-at-bio.umass.edu wrote:
}
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} Greetings,
} I am building a freeze dryer for samples frozen in
} tert-butanol (for SEM -- alternative to critcal point drying). I am
} having difficulty finding information about the resistance of typical
} vacuum pump oils to tert butanol. The pump will be a common rotary
} vane pump and there will be foreline trap. Still, it seems reasonable
} to use an oil that shows good resistance to tBA. Any leads in finding
} one will be gratefully appreciated.
}
} Many thanks!
} Tobias

--
Jacob Kabel
Electron Microscopist
Department of Materials Engineering
The University of British Columbia
6350 Stores Road, Room 419
Vancouver, British Columbia
Canada
V6T 1Z4
Phone: (604) 822-5648
Fax: (604) 822-3619
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Email: frodriguez-at-berkeley.edu
Name: Frances P. Rodriguez-Rivera

Organization: UC Berkeley

Title-Subject: [Filtered] Nanocluster labeling of bacterial cell wall

Message: Hi All,

For the last couple of months I have being trying to label bacterial cell walls (presumably
outermost layer) with electron dense 0.8 nm diameter clusters that should be covalently attached. I
started imaging cluster-labeled bacteria using 400 mesh formvar- and carbon-coated Cu grids using a
Tecnai 12 at 120kV. We were not able to see any labeling.

We carried on the recommendation of imaging our samples using 5nm Si grid (9 windows) with the same
scope. These grids do not have a great affinity for whole-cell bacteria, any recommendations? Also,
at high magnifications (above 100 k) it is very difficult distinguish bacteria (lightly stained with
uranyl acetate) from the grid material. Any suggestions on how to stain bacteria in order to avoid
interference with the nanocluster labeling?

I would greatly appreciate any guidance or suggestions.

Best wishes,
Frances

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Email: rpowell-at-nanoprobes.com
Name: Rick Powell

Organization: Nanoprobes

Title-Subject: [Filtered] Re: TEM - Staining Cells, Nanocluster labeling

Message: Commercial disclosure - we make 0.8 nm gold nanoparticle labeling reagents, as well as
several other reagents mentioned below...

Hello Frances:

You seem to have a couple of different issues going on here.

(1) 0.8 nm gold nanoparticle visibility: it is quite ambitious to expect to easily see 0.8 nm gold
particles in labeled biological specimens by TEM, especially when another electron-dense stain is
present. As Roseann Csencsits mentioned, you should check first whether you can see these particles
on their own on a grid before you try to look for them in labeled specimens.

If you have not already done so, looking at a high-resolution camera image will probably reveal the
particles better than the microscope viewfinder.

If you cannot see them on a grid, then you can either try enlarging them with autometallography -
silver or gold enhancement - or move to larger labels. The 'ultrasmall' 0.8 nm gold labels are about
the same size as the Undecagold (Au11) reagents that we sell (our larger Nanogold reagents, although
not much bigger in terms of diameter at 1.4 nm, contain more than five times the number of gold atoms).

While the ultrasmall 0.8 nm gold labels enhance less readily than some larger gold cluster labels
and nanoparticles, they should do so well enough for you to see them in the TEM. For a couple of
references for this, see:

Flierl, A.; Jackson, C.; Cottrell, B.; Murdock, D.; Seibel, P., and Wallace, D. C.: Targeted
delivery of DNA to the mitochondrial compartment via import sequence-conjugated peptide nucleic
acid. Mol. Ther., 7, 550-557 (2003)

Dellaire, G.; Nisman, R.; Eskiw, C. H., and Bazett-Jones, D. P.: In situ imaging and isolation of
proteins using dsDNA oligonucleotides. Nucleic Acids Res., 32, e165 (2004)

(2) Staining: not sure if you are using uranyl acetate as a negative or positive stain, but you
might consider moving to a stain based on a lower Z element to decrease the electron density
contributed by the stain (we make a negative stain based on vanadium, for example; or ruthenium
instead of osmium). This will obscure the nanoparticle labels less, but you will probably still need
to enhance them.

Hope this helps,

Rick Powell
Nanoprobes, Inc
nanoprobes.com

_____________________________________________________

Hi All,

For the last couple of months I have being trying to label bacterial cell
walls (presumably outermost layer) with electron dense 0.8 nm diameter
clusters that should be covalently attached. I started imaging
cluster-labeled bacteria using 400 mesh formvar- and carbon-coated Cu
grids using a Tecnai 12 at 120kV. We were not able to see any labeling
particles.

Sample prep: Placing sample on glow-discharged grid, let it dry for a few
minutes, and
wick off excess solution.

We carried on the recommendation of imaging our samples using 5 nm Si grid
(9 windows) with the same scope. These grids do not have a great affinity
for whole-cell bacteria, any recommendations? Also, at high magnifications
(above 100 k) it is very difficult distinguish bacteria (lightly stained
with uranyl acetate) from the grid material. Any suggestions on how to
stain bacteria in order to avoid interference with the nanocluster
labeling?

I would greatly appreciate any guidance or suggestions.

Best wishes,
Frances

Frances P. Rodriguez-Rivera
Department of Chemistry
UC Berkeley

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Email: mlibbee-at-gmail.com
Name: Marissa

Organization: LBL

Title-Subject: [Filtered] polymer staining

Message: Hi Listers,

Can anyone recommend a protocol for staining tem thin sections of 70% PVDF 30% acrylate?

Thank you

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Dear colleagues,

I was asked by the Institute of Science and Technology Austria in
Klosterneuburg (in the vicinity of Vienna) to spread the word about a
vacant position for a staff scientist (f/m) in electron microscopy. The
person sought for will lead an EM facility to be established in early
2013, focusing on neuroscience and cell biology.

Details can be found on IST's website:

http://ist.ac.at/about-ist-austria/open-positions/staff-scientists/

Please feel free to forward this message to anyone who might be
interested.

Best regards and have a nice weekend,

Guenter

--
Dr. Guenter Resch
Head of Electron Microscopy, Campus Science Support Facilities GmbH
P: +43 (1) 796 23 24 - 7120; F -227120; W: http://www.csf.ac.at/em
Post: Dr. Bohr-Gasse 3, 1030 Vienna, Austria, European Union

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Wired has a nice featured called "This Day in Tech". Who would have thought that a single lens could be considered high tech? Van Leeuwenhoek and Brian Ford, that's who! By the way, they are not contemporaries and I don't mean to imply they are, I just enjoyed his book.

For more here's a link to Wired.

http://www.wired.com/thisdayintech/2008/09/sept-17-1683-van-leeuwenhoek-gives-us-reason-to-brush-and-floss/

stay safe...........
Frank karl

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Dear Colleagues,

I have an old EM 300 that been experiencing problems with the vacuum
system (for starters). For some time I would activate the automatic
pumping system and the scope would open its main valve in a matter of
5-10 minutes with the HV meter reading zero (I actually doubt it was
active as it never dropped to zero - about 20 was the lowest it ever
got down to). Since I didn't trust the reading I would always wait
another 10-20 minutes before energizing the filament - which lasted
over 80 hours so I had to assume high vacuum was achieved in the
column. The rotary pump would shut down as is typical. These days,
the rotary pump just runs and will not shut off.

I made a visual inspection of the vacuum system and have noted that
the Penning tube is not glowing as in the past (which might explain
my zero reading on the HV meter). I cleaned the tube and tried it
without success. I then located a brand new tube and tried that -
again no success! The previous service engineer stated that he
measured voltage supply to the tube (I have not checked this myself).

Another more recent problem I am experiencing is that I cannot get
the high voltage above 40 kV (during those past rare times when the
vacuum system functioned). The push buttons seem to stick and will
not engage from 60-80-100kV. Even when I can get a button to engage
(for example on 80kV), the voltage meter will only rise to 40 kV. The
image is actually not bad at 40 kV but I am used to running it at
80kV for biological samples.

I am located on Long Island, NY and was wondering if there are any
individuals who can service this vintage TEM in the area. Please
contact me off list if you are a service engineer who can come take a
look at the scope for me. I am running my TEM course this semester
and this is the only TEM I have with a digital camera retrofit.

Thanks for any assistance!

Steve
--
Stephen J. Beck
Professor
Bio-Imaging Center/Electron Microscopy
Department of Biology
Nassau Community College
Garden City, NY 11530
Voice Mail: (516) 572-7829
Email: {becks-at-sunynassau.edu}
URL: {http://www3.ncc.edu/faculty/BIO/becks/BECKS.HTM}

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Title-Subject: [Filtered] cross section

Message: How to cut a 2 micrometer diameter of fiber and make a cross
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Skip Palenik had a method of mounting fibers and hairs between thin sheets of low melting low density polyethylene. The material was clear and you placed the fiber/hair between two sheets of the material, applied a little heat and weight and the plastic fused together. When the sample was cool, (I seem to remember I quenched them in cold water) thin sections were cut under a stereomicroscope with a single edge razor blade.

If you had enough of these fibers, if you can manipulate them, this method may work for you. I'd make several cross sections, mounted them, gold coat them and look with the SEM of the fibers.

Let us know how you solve your problem.

Frank

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Dear listers,

Does anyone have any thoughts on differentiating C14 from C36 laves phases via EBSD? The ICSD reports crystal files for both polytypes of Cr2Zr. I have (Cr,Fe)2Zr EBSD patterns that fit with extremely high confidence to either phase file. Both are the same space group (#194, P63/mmc) so it's no surprise EBSD is confounded. C14 laves is the MgZn2 structure, and C36 is the MgNi2 structure, in which the C36 is more-or-less two C14 unit cells stacked together down c, with a slightly different stacking sequence (http://cst-www.nrl.navy.mil/lattice/struk/ctype.html ).

Looking at predicted band widths (lattice parameters) is less than convincing, due to the similarity of the a-parameters and the fact that the C36 c-parameter is almost exactly twice C14 c-parameter, so high-order reflections tend to fall near the same place. By squinting at the patterns with superimposed predicted bands widths, I think the phase is C14; the literature agrees, but I'd like to convince myself further. Is there anything short of TEM or dynamical simulation I might try?

Thanks,
Chad

PS: Apologies to those of you who also subscribe to the Wisconsin EBSD listserver and have now seen this message twice.

---------------------
Chad M. Parish, Ph.D.
Microscopy Group
Materials Science and Technology Division
Oak Ridge National Laboratory
Phone: 1 865 574 0092
Email: parishcm-at-ornl.gov
Web: www.ornl.gov/share

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Hi Lingling Xu,

I once prepared hair sample cross sections (feline, canine,
human) for SEM with masked broad ion beam milling using a
Gatan Ilion+ 693 (no commercial interest � it�s the only
instrument of its kind that I have used). The hairs were
sandwiched between two 1.7mm thick glass cover slips with
gel super glue, and trimmed close to the edge of the glass
with a scalpel. This sandwich was then mounted on a special
sample �blade� and the hair cross sections ion beam
polished. Finally, I sputter coated with Au/Pd and imaged
with an SEM.

I was able to manipulate my hair samples into place on the
cover slips using a stereoscope. I�m not sure if this would
work with your protein fibers since hair has a much larger
diameter, but it might be worth looking into.

Cheers,

Gigi K.
-----------------------------------

Gigi Kemalyan, Research Technician
HHMI / Nogales Lab -at- UC Berkeley
742 Stanley Hall
Berkeley, CA 94720
Ph. 510-666-3335
cryoem.berkeley.edu

-----------------------------------
No trees were harmed by sending this message; however, a
large number of electrons were greatly inconvenienced.
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Email: xulingling723-at-gmail.com
Name: Lingling Xu

Organization: Dalhousie university

Title-Subject: [Filtered] cross section

Message: How to cut a 2 micrometer diameter of fiber and make a cross
section image by SEM? It is protein fiber, very soft.

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Email: kurganov-at-iptm.ru
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Organization: IMT RAS

Title-Subject: [Filtered] SEM Metal-Coated surface of detectors

Message: Hello everyone!
It's known that the detection surface of scintillator in secondary
electron detectors is coated to prevent charging and for maximum
reflectance. In pioneer ET work thickness of aluminum film was 70 nm.
Today in commercial news-sheet can be read that "Conductive coatings can
be made so thin that they transmit 1keV electrons!" But what is
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Hi

I believe there is some confusion here?

The coating on a scintillator detector is from my experience aluminium
(aluminum) for its low density. As the secondary electrons are only
accelerated by up to 400 volts to bring them into the SE detector I do not
understand the reference to being able to transmit 1Kev electrons? The
only area where I see this may be applied is to backscattered electron
detectors when we are collecting much higher energy electrons with a far
less sensitive detecting system.

Anyone any other ideas?

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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Title-Subject: [Filtered] SEM Metal-Coated surface of detectors

Message: Hello everyone!
It's known that the detection surface of scintillator in secondary electron
detectors is coated to prevent charging and for maximum reflectance. In
pioneer ET work thickness of aluminum film was 70 nm.
Today in commercial news-sheet can be read that "Conductive coatings can be
made so thin that they transmit 1keV electrons!" But what is thickness of
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It should be a typo in the text.
Not 1keV. I mean 1eV.
To my knowledge both BSE and SE scintillation detectors have coatings on
its input surfaces.
Just interesting to know what thickness of this metalized layer in modern
commercial SEMs.

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On Sep 18, 2012, at 10:52 AM, parishcm-at-ornl.gov wrote:

} Does anyone have any thoughts on differentiating C14 from C36 laves
} phases via EBSD? The ICSD reports crystal files for both polytypes
} of Cr2Zr. I have (Cr,Fe)2Zr EBSD patterns that fit with extremely
} high confidence to either phase file. Both are the same space group
} (#194, P63/mmc) so it's no surprise EBSD is confounded. C14 laves is
} the MgZn2 structure, and C36 is the MgNi2 structure, in which the
} C36 is more-or-less two C14 unit cells stacked together down c, with
} a slightly different stacking sequence (http://cst-www.nrl.navy.mil/lattice/struk/ctype.html
} ).
}
} Looking at predicted band widths (lattice parameters) is less than
} convincing, due to the similarity of the a-parameters and the fact
} that the C36 c-parameter is almost exactly twice C14 c-parameter, so
} high-order reflections tend to fall near the same place. By
} squinting at the patterns with superimposed predicted bands widths,
} I think the phase is C14; the literature agrees, but I'd like to
} convince myself further. Is there anything short of TEM or dynamical
} simulation I might try?
}
} Thanks,
} Chad
}
} PS: Apologies to those of you who also subscribe to the Wisconsin
} EBSD listserver and have now seen this message twice.
}
} ---------------------
} Chad M. Parish, Ph.D.
} Microscopy Group
} Materials Science and Technology Division
} Oak Ridge National Laboratory
} Phone: 1 865 574 0092
} Email: parishcm-at-ornl.gov
} Web: www.ornl.gov/share

Dear Chad,
If you can deposit patches of each of the laves onto the specimen--
one hopes they will be epitaxial--EBSD from areas with the appropriate
lave will be completely superimposed. Of course, you would need to
know which of the patches corresponded to which lave and the areas
from which the EBSD data are taken must contain both the unknown and
the known patch. I have no idea whether this is practical for these
substances, so good luck.
Yours,
Bill

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Email: brad-at-nanounity.com
Name: Brad Rangell

Organization: Nanounity

Title-Subject: [Filtered] Tip Enhanced Raman Scattering Workshop at
UPENN In October

Message: NT-MDT will be conducting a Tip Enhanced Raman Scattering
(TERS) workshop hosted by Professor Robert Carpick and the University of
Pennsylvania, October 9, 10 and 11th 2012. TERS is a nanoscale Raman
measurement that requires the integration of AFM and Raman microscopies.
The goal is to achieve sub diffraction optical images and spectra that
correlates to the AFM data.

The workshop will be lead by Dr. Pavel Dorozhkin of NT-MDT and covers
the following subjects:
Live TERS measurement demonstrations!
Lectures on the application of TERS, AFM and Raman in the fields of
physics, chemistry and material science.
Measurements of samples brought by participants of the workshop.

The workshop is free to all participants who register and attend. Space
is limited, so please contact us to ensure your space is reserved!

Dates: October 9,10 and 11th 2012
Location: UPENN Philadelphia, PA

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Brad Rangell
brad-at-nanounity.com
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Folks,
It looks like we'll finally(!) be replacing our MT2B and MT5000. I"m
aware of the Leica UC7 and the RMC/Boekler PT-X(L).
Are there other models out there that I should consider?
Anyone have feedback on the reliability/usability of the models above?

Please respond off-line to smithj-at-winthrop.edu

Thanks in advance,
Julian

--
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Director, Winthrop Microscopy Facility
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803-323-2111 x6427 (vox)
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Dear Collective,

Cleaning out our freezer, I managed to excavate five 250-sheet boxes of Kodak 4489 3 1/4x4 inch sheet film. It's outdated by a few years, but has been frozen continuously.

Does anybody want this film? As I recall it cost way over $1000 to buy this stuff back in the day. Seems a shame to toss it, but toss it we will if there are no takers.

First come, first served!!! The line forms to the left!

Cheers,
Randy

Randy Tindall
Senior Research Specialist
Electron Microscopy Core Facility
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65201
(573) 884-5383

We Do Small Well!

www.emc.missouri.edu (dayjob)
http://nadiasyard.com (blog)
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Hi again,

The film has found a home! If we find more lurking in a corner somewhere, you all will hear from us again.

Thanks all.

Randy

Randy Tindall
Senior Research Specialist
Electron Microscopy Core Facility
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65201
(573) 884-5383

We Do Small Well!

www.emc.missouri.edu (dayjob)
http://nadiasyard.com (blog)
http://www.facebook.com/RandyTindallGalleries (photography)

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Golly! I would not have guessed that Randy's offer of free film would be
taken up so fast. There must still be a wide use of film cameras.

Therefore, I will make my offering to the greater good:

I have 3 unopened 250-sheet boxes of Kodak SO-163, with 2009 date, which are
about to go into the trash. They were kept continually refrigerated until
last week. Yours for the asking.

I am now ducking and covering....

Roger

Roger A. Ristau, PhD
Electron Microscopy Specialist
Institute of Materials Science
97 North Eagleville Road
University of Connecticut
Storrs, CT 06269
vox: 860-486-5453
fax: 860-486-4745
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Thanks for the responses. A home has been found for the SO-163 film.

As I said, there seems to be a large group of electron film user still out
there, so check your closets and make new friends!

Cheers
Roger

Roger A. Ristau, PhD
Electron Microscopy Specialist
Institute of Materials Science
97 North Eagleville Road
University of Connecticut
Storrs, CT 06269
vox: 860-486-5453
fax: 860-486-4745
www.ims.uconn.edu/~micro/

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Anyone have an idea what may be causing out-of-round ring patterns on our
JEOL 2010 LaB6 TEM?

Diffraction patterns on both DualView and MSC cameras show same distortion.
Column alignments were done with care.

I presume there may be contamination (magnetic? I hope not!) on a lens or
otherwise in the column.

Anyone have ideas how to fix it?

Thanks
Roger (my day for thousands of out-of-office replies)

Roger A. Ristau, PhD
Electron Microscopy Specialist
Institute of Materials Science
97 North Eagleville Road
University of Connecticut
Storrs, CT 06269
vox: 860-486-5453
fax: 860-486-4745
www.ims.uconn.edu/~micro/

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Email: acalhoun-at-bidmc.harvard.edu
Name: Andrea Calhoun

Organization: Beth Israel Deaconess Medical Center

Title-Subject: [Filtered] Differences in Mito Contrast/Density

Message: Hello All!

Does anyone know of a comprehensive publication which describes why
mitochondria can display a lighter or darker matrix? Right now I have
cells whose mito are the same density as the cytoplasm, very difficult
to discern. I have had skeletal muscle with intensely dark mito and
liver with light mito. I'm guessing it has something to do with the
protein content?

Its a question I should have found out earlier, alas, time flies
faster than I can keep up with it.

Thank you in advanced for the info!

Best,

^_^ Andrea

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Email: jmircheski [at] us.es
Name: Josif Mircheski

Title-Subject: [Filtered] Re: [Microscopy] purchasing new ultramicrotome

Message: Hi Julian,

I haven't used many different ultramicrotomes, just several of Leica's
models, but I am extremely satisfied with the newest model that our core
facility acquired. It is a Leica UC7 and when I describe it to other
people, I normally say that before, I was driving a Ford Escort, and now
I have a Mercedes. The control is excellent, I've cut sections at 30 nm
without any problem. The control unit has a touch screen where you can
control the lights, the cutting window and some other stuff (different
users, pre-set cutting speeds and thickness, etc.) The thing that I like
the best is that the UC7 that I use comes with a mounted camera under
the binoculars, so you don't have to see through the binoculars all the
time; you can observe the approach on a TV screen, for example. I find
this option excellent especially for teaching purposes, although the
alignment shadows are not always visible on the screen. You can also
record a video of your work with the camera, if you insert a video card.
Also, has two arm pads that are easily adjustable for supporting the
arms/elbows of any user. The UC7 has some other options, but I didn't
need them and can't explain them. The only week point is that the
control of the knife stage (east to west) is still manual. I would love
to see it controlled via the touch screen. In general, I'm extremely
satisfied from this model, as it was a quantum leap from a previous
Rechert Ultracut E.

Disclaimer: Just a very happy user of Leica UC7, no other interest. Not
a user of Ford Escort, nor of any Mercedes (unfortunately).

Best regards from still hot Seville,

Josif

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Email: micro2013-at-canterbury.ac.nz
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Organization: 26th NZ Conference on Microscopy/ MNZ

Title-Subject: [Filtered] Please send out via listserve- thank you

Message: Call for abstracts and registration now open!
26th New Zealand Conference on Microscopy

University of Canterbury, Christchurch, New Zealand
12-15 February 2013

http://www.mech.canterbury.ac.nz/micro2013/

Microscopy New Zealand cordially invites you to their 26th biennial
conference

Hosted by the University of Canterbury this event provides a valuable
opportunity to present research, attend workshops, network with
colleagues, and view the latest equipment and consumables in exhibitor
displays. Keynote speakers include Professor Simon P. Ringer, Professor
David Smith & Dr. Kerry Swanson.

Kicking off with a day of workshops, and continuing into two and a half
days of presentations, the event will also include a welcoming
reception, gala dinner, and an exhibitor & poster reception. Special
registration rates for students are available. Full details on
workshops as well as optional social excursions to be posted online soon.

All enquiries from delegates, sponsors and exhibitors to:
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Josif - that's good to hear as we have just taken delivery of a new UC7!

Julian - yes, when I tendered for one this summer there appeared to be
only 2 manufacturers of ultramicrotomes - Leica and RMC. I had used an
Ultracut E for many (10+) trouble-free years in the pre-digital age; we
did get it annually serviced though. For this and other reasons I
decided to buy the UC7. I haven't used the RMC XT but it looks like a
good machine too (check user comments on this listserver), and was
competitively priced.

Regards
Mark

Dr Mark A.E. Auty
National Food Imaging Centre
Food Chemistry & Technology Department
Teagasc Food Research Centre
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Fermoy, Co. Cork, Ireland

tel +353 25 42442
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Email: jmircheski [at] us.es
Name: Josif Mircheski

Title-Subject: [Filtered] Re: [Microscopy] purchasing new ultramicrotome

Message: Hi Julian,

I haven't used many different ultramicrotomes, just several of Leica's
models, but I am extremely satisfied with the newest model that our core

facility acquired. It is a Leica UC7 and when I describe it to other
people, I normally say that before, I was driving a Ford Escort, and now

I have a Mercedes. The control is excellent, I've cut sections at 30 nm
without any problem. The control unit has a touch screen where you can
control the lights, the cutting window and some other stuff (different
users, pre-set cutting speeds and thickness, etc.) The thing that I like

the best is that the UC7 that I use comes with a mounted camera under
the binoculars, so you don't have to see through the binoculars all the
time; you can observe the approach on a TV screen, for example. I find
this option excellent especially for teaching purposes, although the
alignment shadows are not always visible on the screen. You can also
record a video of your work with the camera, if you insert a video card.

Also, has two arm pads that are easily adjustable for supporting the
arms/elbows of any user. The UC7 has some other options, but I didn't
need them and can't explain them. The only week point is that the
control of the knife stage (east to west) is still manual. I would love
to see it controlled via the touch screen. In general, I'm extremely
satisfied from this model, as it was a quantum leap from a previous
Rechert Ultracut E.

Disclaimer: Just a very happy user of Leica UC7, no other interest. Not
a user of Ford Escort, nor of any Mercedes (unfortunately).

Best regards from still hot Seville,

Josif

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Hi

The most common reason I have seen for a distortion in the diffraction
pattern is intermediate astigmatism. On the other hand if you have a
specimen with a regular structure, a good example is a cross grating carbon
replica, do you notice that this image is distorted too? Magnetic fields or
vibration would blur the image in one direction, visible as a softening 180
degs apart.

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: raristau-at-ims.uconn.edu [mailto:raristau-at-ims.uconn.edu]
Sent: 20 September 2012 18:45
To: protrain-at-emcourses.com

Anyone have an idea what may be causing out-of-round ring patterns on our
JEOL 2010 LaB6 TEM?

Diffraction patterns on both DualView and MSC cameras show same distortion.
Column alignments were done with care.

I presume there may be contamination (magnetic? I hope not!) on a lens or
otherwise in the column.

Anyone have ideas how to fix it?

Thanks
Roger (my day for thousands of out-of-office replies)

Roger A. Ristau, PhD
Electron Microscopy Specialist
Institute of Materials Science
97 North Eagleville Road
University of Connecticut
Storrs, CT 06269
vox: 860-486-5453
fax: 860-486-4745
www.ims.uconn.edu/~micro/

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Steve,
The typical secondary electron impacts the scintillator with an energy in
the range of 8-12keV, due to the voltage applied to the scintillator
directly. This is why a secondary electron detector can be used as a
backscattered electron detector by turning off the scintillator high voltage
as opposed to making the Faraday Cage voltage negative. Either way works.
The first makes secondaries unable to stimulate the phosphor, the second
keeps secondaries from even impacting the scintillator.

M. E. Taylor Engineering has been making scintillators without any Al
coating for at least 2 decades by using a conductive binder for the
phosphor. This gives greater sensitivity but doesn't assist in reflecting
light back down the pipe, but I don't feel that is very important given how
well Gene's scintillators work. I recommend his scintillators to my
customers but have not had a wholesale relationship for quite a few years.

As for Iliya's thoughts about coating thickness, the conductive binder makes
that moot. However, I would think that putting a coating thin enough for a
1keV (or as later stated 1eV) electron to penetrate would 1) prove so
fragile as to be a non-conductive coating after just a few vacuum/humidity
cycles and 2)not gain you much of anything because I'm not sure a 1keV (let
alone a 1eV) electron could excite the phosphor anyway. I've always noticed
that even backscattered electrons are pretty much undetectable below a 5kV
beam voltage with most detectors (including Everly-Thornhart detectors with
no voltage on the scintillator). An E-T detector with +10kV on the
scintillator and -400V on the Faraday cage would be an exception because the
BSEs would be further accelerated by the scintillator voltage and would
light up the phosphor.

I think it requires a micro-channel detector to effectively detect
secondaries without serious acceleration, or the gas phase detectors used in
some ESEMs for high pressure work.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: protrain-at-emcourses.com [mailto:protrain-at-emcourses.com]
Sent: Wednesday, September 19, 2012 10:01 AM
To: kenconverse-at-qualityimages.biz

Hi

I believe there is some confusion here?

The coating on a scintillator detector is from my experience aluminium
(aluminum) for its low density. As the secondary electrons are only
accelerated by up to 400 volts to bring them into the SE detector I do not
understand the reference to being able to transmit 1Kev electrons? The
only area where I see this may be applied is to backscattered electron
detectors when we are collecting much higher energy electrons with a far
less sensitive detecting system.

Anyone any other ideas?

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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To: protrain-at-emcourses.com

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Email: kurganov-at-iptm.ru
Name: Iliya Kurganov

Organization: IMT RAS

Title-Subject: [Filtered] SEM Metal-Coated surface of detectors

Message: Hello everyone!
It's known that the detection surface of scintillator in secondary electron
detectors is coated to prevent charging and for maximum reflectance. In
pioneer ET work thickness of aluminum film was 70 nm.
Today in commercial news-sheet can be read that "Conductive coatings can be
made so thin that they transmit 1keV electrons!" But what is thickness of
conductive films in modern SEMs detectors? Can anyone give examples of
coating thickness for detector in any modern SEM?

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Email: achristensen-at-annamaria.edu
Name: Arne Christensen

Organization: Anna Maria College

Title-Subject: [Filtered] Advice/support for a microscopist

Message: Dear listeners,

Im a new faculty member at a small liberal arts school, and like many
of us, Im working within the confines of a diminishing budget and
resources. In my previous professional lives Ive been an avid user of
various types of microscopy, and incorporated their use in my courses
and research. In my present position I am limited in this respect, as
our catalog of microscopes is limited to low-end standard teaching
microscopes. Because my institution is primarily a teaching college,
securing a large equipment grant or research grant is not an option.

Im trying a couple avenues to try and acquire a florescence microscope;
seeking equipment donations, used equipment at a discounted price,
and/or a grant to purchase equipment for use in the classroom. Im
reaching out to this community to see if anyone can advise me on any of
these fronts. If anyone is aware of equipment for donation, it is
possible I could request support from my administration for
packing/shipping or depending on location I could do it myself.

Here is some course work I conducted using microscopy in the classroom
at my previous position:

http://web.york.cuny.edu/~achristensen/DevBio.html

I hope this listserver is an appropriate place to post this message,
Ive been a listener for some time, and I seem to recall similar
messages in the past.

Thanks,
AC

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Hello All,

We are pleased to announce that on this coming Friday, September 28th, we will be having the�Hans Ris Symposium on 3D Electron Microscopy. This symposium is in honor of Hans Ris, one of the giants in cell biology and electron microscopy, who was a Professor at the UW from 1949-2004. The event is free to all (no registration) and will overview the latest in biological electron microscopy applications, include cyro EM and Electron Tomography.

The complete schedule and other information is here:�http://www.microscopy.wisc.edu/events/hansrissymposium2012

Please let me know if you have any questions.

best
kevin

--
Kevin W. Eliceiri
Director,�Laboratory for Optical and Computational Instrumentation (LOCI)
Departments Cell and Molecular Biology and Biomedical Engineering
Affiliate Principal Investigator, Morgridge Institute for Research (MIR)
Room 271 Animal Sciences,�1675 Observatory Drive,�Madison, WI 53706
Phone: 608-263-6288

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Hi Ken

I fully understand what you are describing. I was just puzzled at the
thought of attracting 1keV electrons into an E-T detector when the
attractive force is no more than 400v!

In answer to low voltage BSE, some of the latest units have been good enough
for me to use them at 2.5kV, made possible through a very thin surface
coating; gold I believe.

Regards

Steve

-----Original Message-----
X-from: kenconverse-at-qualityimages.biz [mailto:kenconverse-at-qualityimages.biz]
Sent: 22 September 2012 02:08
To: protrain-at-emcourses.com

Steve,
The typical secondary electron impacts the scintillator with an energy in
the range of 8-12keV, due to the voltage applied to the scintillator
directly. This is why a secondary electron detector can be used as a
backscattered electron detector by turning off the scintillator high voltage
as opposed to making the Faraday Cage voltage negative. Either way works.
The first makes secondaries unable to stimulate the phosphor, the second
keeps secondaries from even impacting the scintillator.

M. E. Taylor Engineering has been making scintillators without any Al
coating for at least 2 decades by using a conductive binder for the
phosphor. This gives greater sensitivity but doesn't assist in reflecting
light back down the pipe, but I don't feel that is very important given how
well Gene's scintillators work. I recommend his scintillators to my
customers but have not had a wholesale relationship for quite a few years.

As for Iliya's thoughts about coating thickness, the conductive binder makes
that moot. However, I would think that putting a coating thin enough for a
1keV (or as later stated 1eV) electron to penetrate would 1) prove so
fragile as to be a non-conductive coating after just a few vacuum/humidity
cycles and 2)not gain you much of anything because I'm not sure a 1keV (let
alone a 1eV) electron could excite the phosphor anyway. I've always noticed
that even backscattered electrons are pretty much undetectable below a 5kV
beam voltage with most detectors (including Everly-Thornhart detectors with
no voltage on the scintillator). An E-T detector with +10kV on the
scintillator and -400V on the Faraday cage would be an exception because the
BSEs would be further accelerated by the scintillator voltage and would
light up the phosphor.

I think it requires a micro-channel detector to effectively detect
secondaries without serious acceleration, or the gas phase detectors used in
some ESEMs for high pressure work.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: protrain-at-emcourses.com [mailto:protrain-at-emcourses.com]
Sent: Wednesday, September 19, 2012 10:01 AM
To: kenconverse-at-qualityimages.biz

Hi

I believe there is some confusion here?

The coating on a scintillator detector is from my experience aluminium
(aluminum) for its low density. As the secondary electrons are only
accelerated by up to 400 volts to bring them into the SE detector I do not
understand the reference to being able to transmit 1Kev electrons? The
only area where I see this may be applied is to backscattered electron
detectors when we are collecting much higher energy electrons with a far
less sensitive detecting system.

Anyone any other ideas?

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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Sent: 19 September 2012 12:49
To: protrain-at-emcourses.com

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Email: kurganov-at-iptm.ru
Name: Iliya Kurganov

Organization: IMT RAS

Title-Subject: [Filtered] SEM Metal-Coated surface of detectors

Message: Hello everyone!
It's known that the detection surface of scintillator in secondary electron
detectors is coated to prevent charging and for maximum reflectance. In
pioneer ET work thickness of aluminum film was 70 nm.
Today in commercial news-sheet can be read that "Conductive coatings can be
made so thin that they transmit 1keV electrons!" But what is thickness of
conductive films in modern SEMs detectors? Can anyone give examples of
coating thickness for detector in any modern SEM?

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Email: amit.welcomes.u-at-gmail.com
Name: Amit

Organization: indian institute of science

Title-Subject: [Filtered] TEM correcting beam tilt (?)

Message: Thank you every one who replied now a major problem in solved.
I aligned gun shift and tilt more meticulously and avoided too excited
C2 so those streaks dont appear as of now. But one more thing keeps
bothering me still is the unsymmetry on beam.
I took 2 more pics of beam as shown in the link below (bottom 2)we can
see such "spikes" on one half of the beam.
What are they?
anything that degrades the quality?
Any way to correct them?

Also i noticed that instead of being completely circular beam is bit
"flat" on one side, like the shape of partially deflated football. any
suggestion/ explanations/ guesses are welcome.

Thanks in advance

https://sites.google.com/site/auxilliarylinks/

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Hi Amit,

The "spikes" are due to a dirty beam-defining aperture (usually the C2
aperture). Something on the aperture edge is charging. Replace or clean
the aperture and you should be good to go!

As a quick test, you should be able to change to a different C2 aperture
and if that aperture is clean, you should see an improved beam. Also be
sure to do a proper C2 aperture centering.

Cheers,
Henk

At 9/22/2012 4:57 PM, microscopylistserver-noreply-at-microscopy.com wrote:
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}
} Message: Thank you every one who replied now a major problem in solved.
} I aligned gun shift and tilt more meticulously and avoided too excited
} C2 so those streaks dont appear as of now. But one more thing keeps
} bothering me still is the unsymmetry on beam.
} I took 2 more pics of beam as shown in the link below (bottom 2)we can
} see such "spikes" on one half of the beam.
} What are they?
} anything that degrades the quality?
} Any way to correct them?
}
} Also i noticed that instead of being completely circular beam is bit
} "flat" on one side, like the shape of partially deflated football. any
} suggestion/ explanations/ guesses are welcome.
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
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"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

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Hello Amit,
your third TEM image with the "spikes" for me has the look of a dirty inliner surface / dirty aperture hole causing astigmatismn
on the beam.
Did you try other (contrast) apertures and did you do a careful alignment of the condensor astigmatism?
There seems also to be an orientation pattern in the edge of the structure. Did you use a mesh of a hexagonal grid when shooting
the image?

Best wishes,
Stefan

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

Am 22.09.12 23:00, schrieb microscopylistserver-noreply-at-microscopy.com:
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} Name: Amit
}
} Organization: indian institute of science
}
} Title-Subject: [Filtered] TEM correcting beam tilt (?)
}
} Message: Thank you every one who replied now a major problem in solved.
} I aligned gun shift and tilt more meticulously and avoided too excited
} C2 so those streaks dont appear as of now. But one more thing keeps
} bothering me still is the unsymmetry on beam.
} I took 2 more pics of beam as shown in the link below (bottom 2)we can
} see such "spikes" on one half of the beam.
} What are they?
} anything that degrades the quality?
} Any way to correct them?
}
} Also i noticed that instead of being completely circular beam is bit
} "flat" on one side, like the shape of partially deflated football. any
} suggestion/ explanations/ guesses are welcome.
}
} Thanks in advance
}
} https://sites.google.com/site/auxilliarylinks/
}
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9, 22 -- From stefan.diller-at-t-online.de Sun Sep 23 04:12:25 2012
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Thank you all for replies, Stefan- no i was not using any hexagonal
grids, and these spikes were there despite careful alignment. So today
i used different condenser aperture and beam was as expected,
symmetrical. So its a dirty aperture only. Any way to correct it my
self? or only removing it is the way?

one more question, look at the 2nd pic in line, the one i posted
earlier, there we can see one side of beam bit different than other
(darkening more) can it also be attributed to the same?

Thanks again

On 9/23/12, stefan.diller-at-t-online.de {stefan.diller-at-t-online.de} wrote:
}
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}
} Hello Amit,
} your third TEM image with the "spikes" for me has the look of a dirty
} inliner surface / dirty aperture hole causing astigmatismn
} on the beam.
} Did you try other (contrast) apertures and did you do a careful alignment of
} the condensor astigmatism?
} There seems also to be an orientation pattern in the edge of the structure.
} Did you use a mesh of a hexagonal grid when shooting
} the image?
}
} Best wishes,
} Stefan
}
}
}
}
} -----------------------------------------------------
} Stefan Diller - Scientific Photography
} Arndtstrasse 22
} D - 97072 Wuerzburg Germany
} ++49-931-7848700 Phone
} ++49-931-7848701 Fax
} ++49-175-7177051 Mobile
}
} Websites:
} www.stefan-diller.com
} www.electronmicroscopy.info
} www.elektronenmikroskopie.info
} www.assisi.de
} www.zwillingsprojekt.de
} Anfahrt: http://Mail.map24.com/Stefan.Diller
} -----------------------------------------------------
}
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} } Email: amit.welcomes.u-at-gmail.com
} } Name: Amit
} }
} } Organization: indian institute of science
} }
} } Title-Subject: [Filtered] TEM correcting beam tilt (?)
} }
} } Message: Thank you every one who replied now a major problem in solved.
} } I aligned gun shift and tilt more meticulously and avoided too excited
} } C2 so those streaks dont appear as of now. But one more thing keeps
} } bothering me still is the unsymmetry on beam.
} } I took 2 more pics of beam as shown in the link below (bottom 2)we can
} } see such "spikes" on one half of the beam.
} } What are they?
} } anything that degrades the quality?
} } Any way to correct them?
} }
} } Also i noticed that instead of being completely circular beam is bit
} } "flat" on one side, like the shape of partially deflated football. any
} } suggestion/ explanations/ guesses are welcome.
} }
} } Thanks in advance
} }
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--
-With Regards
Amit Gupta
Research Scholar
IISc

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Steve,
Sorry I missed that angle. Yes, the Faraday cage might have some difficulty
drawing 1keV electrons towards the detector. Are the detectors that work at
2.5kV diode or phosphor/light pipe?
Ken

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
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qualityimages.biz

-----Original Message-----
X-from: Steve Chapman [mailto:protrain-at-emcourses.com]
Sent: Saturday, September 22, 2012 2:59 PM
To: kenconverse-at-qualityimages.biz
Cc: Microscopical Soc of America

Steve,
The typical secondary electron impacts the scintillator with an energy in
the range of 8-12keV, due to the voltage applied to the scintillator
directly. This is why a secondary electron detector can be used as a
backscattered electron detector by turning off the scintillator high voltage
as opposed to making the Faraday Cage voltage negative. Either way works.
The first makes secondaries unable to stimulate the phosphor, the second
keeps secondaries from even impacting the scintillator.

M. E. Taylor Engineering has been making scintillators without any Al
coating for at least 2 decades by using a conductive binder for the
phosphor. This gives greater sensitivity but doesn't assist in reflecting
light back down the pipe, but I don't feel that is very important given how
well Gene's scintillators work. I recommend his scintillators to my
customers but have not had a wholesale relationship for quite a few years.

As for Iliya's thoughts about coating thickness, the conductive binder makes
that moot. However, I would think that putting a coating thin enough for a
1keV (or as later stated 1eV) electron to penetrate would 1) prove so
fragile as to be a non-conductive coating after just a few vacuum/humidity
cycles and 2)not gain you much of anything because I'm not sure a 1keV (let
alone a 1eV) electron could excite the phosphor anyway. I've always noticed
that even backscattered electrons are pretty much undetectable below a 5kV
beam voltage with most detectors (including Everly-Thornhart detectors with
no voltage on the scintillator). An E-T detector with +10kV on the
scintillator and -400V on the Faraday cage would be an exception because the
BSEs would be further accelerated by the scintillator voltage and would
light up the phosphor.

I think it requires a micro-channel detector to effectively detect
secondaries without serious acceleration, or the gas phase detectors used in
some ESEMs for high pressure work.

Ken Converse
owner

QUALITY IMAGES
Servicing Scanning Electron Microscopes
Since 1981
474 So. Bridgton Rd.
Bridgton, ME 04009
207-647-4348
kenconverse-at-qualityimages.biz
qualityimages.biz

-----Original Message-----
X-from: protrain-at-emcourses.com [mailto:protrain-at-emcourses.com]
Sent: Wednesday, September 19, 2012 10:01 AM
To: kenconverse-at-qualityimages.biz

Hi

I believe there is some confusion here?

The coating on a scintillator detector is from my experience aluminium
(aluminum) for its low density. As the secondary electrons are only
accelerated by up to 400 volts to bring them into the SE detector I do not
understand the reference to being able to transmit 1Kev electrons? The
only area where I see this may be applied is to backscattered electron
detectors when we are collecting much higher energy electrons with a far
less sensitive detecting system.

Anyone any other ideas?

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

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Email: kurganov-at-iptm.ru
Name: Iliya Kurganov

Organization: IMT RAS

Title-Subject: [Filtered] SEM Metal-Coated surface of detectors

Message: Hello everyone!
It's known that the detection surface of scintillator in secondary electron
detectors is coated to prevent charging and for maximum reflectance. In
pioneer ET work thickness of aluminum film was 70 nm.
Today in commercial news-sheet can be read that "Conductive coatings can be
made so thin that they transmit 1keV electrons!" But what is thickness of
conductive films in modern SEMs detectors? Can anyone give examples of
coating thickness for detector in any modern SEM?

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Hi Amit,

Glad the aperture problem is diagnosed. Apertures can be cleaned, check
your microscope owners guide for the procedure. It will differ
depending on whether you are using Pt or Mo apertures. The other option
is to just replace the dirty aperture.

The uneven illumination in the 2nd photo *appears* to be due to the "hot
spot" in the emission of a FEG source. I assume that your microscope
has a Schottky "FEG" emitter. The emission from a Schottky source is
strongly distributed in the forward direction and falls off at larger
emission angles. If you use a large beam limiting aperture (usually C2)
you can see this distribution (so-called "witches hat profile"). For my
Tecnai F20, I see something like this when I use a 150um C2 aperture.
The fact that the hot spot is off center means that the gun alignment is
off by a little bit. This is corrected by using the gun tilt and shift.
Once you get the "hot spot" centered in the large aperture, you will
generally operate with a smaller C2 aperture which cuts off the less
intense "halo" around the central "hot spot".

Good luck,
Henk

At 9/23/2012 6:18 AM, amit.welcomes.u-at-gmail.com wrote:
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} Thank you all for replies, Stefan- no i was not using any hexagonal
} grids, and these spikes were there despite careful alignment. So today
} i used different condenser aperture and beam was as expected,
} symmetrical. So its a dirty aperture only. Any way to correct it my
} self? or only removing it is the way?
}
} one more question, look at the 2nd pic in line, the one i posted
} earlier, there we can see one side of beam bit different than other
} (darkening more) can it also be attributed to the same?
}
} Thanks again
}
}
} On 9/23/12, stefan.diller-at-t-online.de {stefan.diller-at-t-online.de} wrote:
} }
} }
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} }
} } Hello Amit,
} } your third TEM image with the "spikes" for me has the look of a dirty
} } inliner surface / dirty aperture hole causing astigmatism
} } on the beam.
} } Did you try other (contrast) apertures and did you do a careful alignment of
} } the condensor astigmatism?
} } There seems also to be an orientation pattern in the edge of the structure.
} } Did you use a mesh of a hexagonal grid when shooting
} } the image?
} }
} } Best wishes,
} } Stefan
} }
} }
} }
} }
} } -----------------------------------------------------
} } Stefan Diller - Scientific Photography
} } Arndtstrasse 22
} } D - 97072 Wuerzburg Germany
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} } } Email: amit.welcomes.u-at-gmail.com
} } } Name: Amit
} } }
} } } Organization: indian institute of science
} } }
} } } Title-Subject: [Filtered] TEM correcting beam tilt (?)
} } }
} } } Message: Thank you every one who replied now a major problem in solved.
} } } I aligned gun shift and tilt more meticulously and avoided too excited
} } } C2 so those streaks dont appear as of now. But one more thing keeps
} } } bothering me still is the unsymmetry on beam.
} } } I took 2 more pics of beam as shown in the link below (bottom 2)we can
} } } see such "spikes" on one half of the beam.
} } } What are they?
} } } anything that degrades the quality?
} } } Any way to correct them?
} } }
} } } Also i noticed that instead of being completely circular beam is bit
} } } "flat" on one side, like the shape of partially deflated football. any
} } } suggestion/ explanations/ guesses are welcome.
} } }
} } } Thanks in advance
} } }
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

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Hi Mauricio,

Generally, refill can be done by anyone who is capable of re-installing
and aligning the GIS after refill: XeF2 powder is available in small
quantities from Praxair, Alfa Aesar, or Strem. Use fume hood, gloves,
gown if you are superstitious, glasses or face mask, and make sure that
XeF2 powder has absolutely no contact with moisture or anything organic
- other then these and other basic precautions commonly used while
handling corrosive and poisonous materials the refill is a very
straight-forward operation. I always use five-nines (99.999%) grade
XeF2, but know of people who are using 99.5% variety for cost reasons
and claim to have no complains.

Note that (depending on OEM and design version of your GIS) there may be
a small O-ring in the central channel of the crucible - make sure to not
get any XeF2 powder on it. It is a good idea to replace or at least
inspect this O-ring every refill, or at least every other refill.
Removing the O-ring is a bit tricky, unless you make a plastic tool for
it. I use Kalrez, but have no idea what kind of material is OEM using there.

If you prefer to not do the refill on your own then feel free to contact
me off the list and I'd be happy to give you a quote.

Important disclaimer: the information above is provided "for educational
and entertainment purposes only." Make sure that you are trained,
qualified, and authorized to handle corrosive and poisonous chemicals at
your institution.

Happy FIBing :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 9/24/2012 1:08 PM, oshel1pe-at-cmich.edu wrote:
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} Forwarded from "Ask a Microscopist"
} Please remember that the person asking the question is likely
} not a member the listserver, and
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} as well as to the list.
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} to the person asking the question.
} Please copy their email address from their question.
} ****************************************************************************************
} } realname - Mauricio Bravo
} } Email - mauricio.bravo-at-maximintegrated.com
} } EDUCATION - Graduate College
} } LOCATION - Dallas, Texas, USA
} } SUBJECT_OF_QUESTION - FIB Gas Supplies - Xenon difluoride GIS Refill
} } QUESTION - Hi!
} } I need to refill my Insulator Enhansed Etch (IEE, Xenon DIfluoride)
} } Gas Injection System (GIS), can you recommend a company or
} } lavoratory that provides that service?
} }
} } Many thanks in advance for your time.
} }
} } Best regards,
} } Mauricio
}
}

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8, 38 -- GIS Refill
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I have a colleague looking for

an EDAX DX-4 computer tower used with EDS systems circa '90s

If you know of any available, please email me.

Tony

.........................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
Environmental, Health & Safety, Microscopy, & Forensic Engineering
pH2, LLC
5250 E US Highway 36, Suite 830
Avon, IN 46123
(317) 718-7020 Office
(317) 718-7038 Fax
(317) 409-3238 Cell
aahavics-at-pH2LLC.com
www.ph2LLC.com

==============================Original Headers==============================
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Hello,
I need a copy (PDF) of the two conference papers at�MICROSCOPY AND MICROANALYSIS 1998 and 1999. We have an access to the E-journal, but I cannot find Supplement.�Our library is still finding a potential lender for more than two weeks. I appreciate it if you could provide a copy of the following proceedings. Thank you for your help.

Hiromi Konishi
UW-Madison
--------------------------------------

Article title
In situ TEM study of inert fillers in liquid environment
Author
Chiou, W.-A. Lee, Y.-C. Ishikawa, A. Konishi, H. Fukushima, K. Shriver, D. F.
Journal title
MICROSCOPY AND MICROANALYSIS
Bibliographic details 1998, VOL 4; SUPP/2, pages 820-821

Article title
In Situ TEM Study Of DNA/Gold Nanoparticles In Liquid Environment
Author
Chiou, W. A. Mucic, R. Ishikawa, A. Konishi, H. Fukushima, K. Mirkin, C.
Journal title
MICROSCOPY AND MICROANALYSIS -NEW YORK-
Bibliographic details 1999, VOL 5; SUPP/2, pages 340-341

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Hello,

I received a copy of the proceedings. Thank you for your help.

Hiromi Konishi
UW-Madison

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Greetings,

I know this is a bit of a loaded question, but is there anyone out
there with a service manual for a JEOL JSM-6XXXF Field emission
service manual? Anything in the 6000 series should work for me- 6300,
6400, etc. I have the schematics and regular operating manual, but I
really need a service manual. Please contact me off-list.

Any help would be greatly appreciated.

--Justin A. Kraft

--
"America believes in education; the average professor earns more money
in a year than a professional athlete earns in a whole week." Evan
Esar

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6, 26 -- Subject: Field service guide?
6, 26 -- From: Justin Kraft {kraftpiano-at-gmail.com}
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6, 26 -- Content-Type: text/plain; charset=ISO-8859-1
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Title-Subject: [Filtered] FEI Q200 Spare Parts

Message: Hi,

I am looking for a spare parts source for FEI MK1 SEM.

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Name: Vince Crist

Organization: Nanolab Technologies

Title-Subject: [Filtered] D-SIMS used systems - want to buy

Message: If you have a used D-SIMS system, please write to us. We are
very interested to buy 2 used D-SIMS systems.
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Greetings Fellow Microscopists,

A joint meeting of M3S and the EM program at Madison College (formerly Madison Area Technical College), will be held in Madison, WI, on Friday, October 19, 2012. This will be a vendor symposium featuring new and breakthrough microscopy instrumentation and products. Program details and registration information can be found on the M3S website Meetings page:

http://www.midwestmicroscopy.org/

Please contact Michael Kostrna at 608-243-4309, or mkostrna-at-madisoncollege.edu with inquiries about the meeting. We look forward to seeing you in Madison!

Elaine Schumacher
M3S Program Coordinator

*********************************************************************
Elaine F.Schumacher
Senior Research Scientist
McCrone Associates, Inc.
850 Pasquinelli Drive
Westmont, IL 60559-5539 USA
630-887-7100 (tel)
630-887-7417 (fax)
E-mail: eschumacher-at-mccrone.com
Web Site: www.mccrone.com

*********************************************************************
This message and any attachments are solely for the
intended recipient. If you are not the intended recipient,
disclosure, copying, use or distribution of the information
included in this message is prohibited.
*********************************************************************

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12, 28 -- Date: Tue, 25 Sep 2012 14:11:40 -0500
12, 28 -- Subject: Meeting Announcement: Madison College and Midwest Microscopy and
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Dear Listers, wherever you may be located:
good morning, good afternoon, good evening everywhere !

The Society for Cutaneous Ultrastructure Research (SCUR) invites you to
participate in its
-------------------
40th Annual meeting which at the same time will be held as the
6th Joint Meeting with the (Japanese) Society for Skin Structure Research
-------------------
taking place in
SALZBURG, AUSTRIA from Sunday, May 12th - Tuesday May 14th, 2013.
(this meeting will take place right after the IID-Meeting in Edinburgh, UK)

General Topic: "From Genetics to Therapy"
The Board Members of the SCUR and the SSSR as well as the Organizing Committee are looking forward to welcoming YOU in Salzburg in May 2013.
Our Annual Conferences and Joint Meetings are known as familial meetings which are well organized, promising highlights - scientifically as well as culturally - and permit for direct exchanges between renowned specialists in the many fields of ultrastructural and molecular cutaneous research.

Invited Key Note Speakers so far:

- Prof. Dr. Hideya ANDO, Okayama (Japan)
[proposed topic: "Electron Microscopic Observation of Melanosome Transfer from Melanocytes to Keratinocytes and Fibroblasts"]

- Prof. Dr. Masamitsu ICHIHASHI, Doshisha University (Japan)
[proposed topic: "Anti-Aging Medicine in Dermatology"]

- Prof. Dr. David KELSELL, London (UK)
[proposed topic: "Connexins & Another Junctional Protein related to Skin Disease"]

- Prof. Dr. John McGRATH, London (UK)
[proposed topic: "Value of Electron Microscopy in today�s world of Molecular Diagnostics"]

- PD Dr. Dr. Angelika RIEMER, Heidelberg (D)
[proposed topic: "HPV and Immunotherapies"]

- Prof. Dr. Nikolaus ROMANI, Innsbruck (A)
[proposed topic: "The Langerhans Cell - History and Modern Research"]

---------------------------------
SCUR & SSSR 2013 SALZBURG/Austria
---------------------------------
Organizing Committee:
Prof. Dr. Helmut Hintner, Prof. Dr. Johann Bauer, Dr. Wolfgang Muss, and Dr. Rudolf Hametner
Department of Dermatology and
Institute of Pathology, General Hospital Salzburg
and Paracelsus Medical University Salzburg, in
Cooperation with the Study Group for Genetics & Regenerative Medicine of the Austrian Society for Dermatology and Venerology

For the First Announcement visit the Website of SCUR:
http://www.scur.org (go to Link {Next Meeting} )
or inquire your personal pdf's via e-mail from the
Secretary of SCUR: Dr. Wolfgang MUSS
( w.muss-at-salk.at or scursssr.salzbg2013-at-Gmail.com )

If you would like to get further information on the Conference by e-mail (list of interested colleagues) please also just send your request to the Secretary.
========================================================
The meeting location (and host hotel) is the

CASTELLANI Parkhotel Salzburg {modern meets classic}
========================================================
situated only 5-10 minutes walk from the very Old City Center of SALZBURG
and easily can be reached by public busses or Taxi from the Main Railway Station (approx. 5 km/10-15 min) and the Airport W. A. Mozart (approx. 7 km/10-15 min). For our guests / participants we have made options for accommodation at a special reduced rate.

All necessary information (Call for Papers, Abstract Deadline, Registration, Accommodation, Social Program, etc.) will be circulated end of October 2012

We hope to see you there!

Best regards,

Wolfgang MUSS PhD
Secretary of SCUR
Member of MSA

==============================Original Headers==============================
24, 37 -- From W.Muss-at-salk.at Wed Sep 26 05:31:28 2012
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Name: See Wee Chee

Organization: Rensselaer Polytechnic Institute

Title-Subject: [Filtered] Digital Micrograph Menu Bar Bug

Message: Hi! I have an issue with my active windows in GIF control
getting stuck underneath the menu bar and I cannot pull the windows out.

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Email: erwrigh-at-emory.edu
Name: Elizabeth Wright

Organization: Emory University

Title-Subject: [Filtered] POSTDOCTORAL FELLOW POSITIONS, EMORY UNIVERSITY

Message: Postdoctoral fellow positions are available in the Wright
laboratory at Emory University starting fall/winter 2012
(http://electronmicroscopy.emory.edu). Research in the lab is focused on
bacterial and viral structural biology, with emphasis on 3-D
structure/function relationships between host cells and viruses. Areas
of interest include cryo-electron tomography of bacteria, bacteria
bacteriophage interactions, paramyxoviruses, and HIV-1. Technology
development interests are in the areas of correlative microscopy and
phase plate cryo-microscopy.

The first postdoctoral position is aimed at studies of the structure of
flagella and type IV pili from several bacterial species. The long-term
goal of the project is to understand 1) the structural implications of
the incorporation of multiple flagellins into the flagellar filament and
2) the structural variation between T4bP and Flp-type pili at the
macromolecular level.

The second postdoctoral position is to develop of correlative microscopy
(fluorescent to cryo-EM) techniques and novel macromolecular probes to
study aspects of viral fusion, entry, assembly, and trafficking. The
viral systems to be studied include HIV-1 and several members of the
paramyxovirus family.

Candidates should have an interest in structural biology, bacteriology
or virology, and should enjoy working as part of an active research
team. Background experience in one (or more) of the following fields is
desirable, but not required: bacteriology, virology, (cryo-) electron
microscopy, and/or image processing/analysis techniques.

The wet laboratory is within the Division of Pediatric Infectious
Diseases and is fully equipped for all aspects of microbiology,
molecular biology, molecular virology, and protein biochemistry. The
Wright lab has complete access to a JEOL 2200FS 200 kV FEG TEM with an
in-column energy filter, Zernike-style phase plates, and high-resolution
4kx4k CCD camera; a JEOL 1400 120 kV TEM with 2kx2k camera; an FEI
Vitrobot and a manual grid plunge freezer; a Bal-Tec high pressure
freezer and Leica freeze substitution system; and a Leica
cryo-ultramicrotome. Emory is a vibrant, multidisciplinary campus with
strong ties to Georgia Tech and other universities within Georgia.
Access to facilities in NMR, Mass Spectrometry, and X-ray
crystallography are available.

Applicants should include a cover letter describing research experience
and interests, curriculum vitae, and names and contact information for
three references.

Dr. Elizabeth R. Wright
Emory University
Department of Pediatrics
2015 Uppergate Drive, NE Room: 548
Atlanta, GA 30322
erwrigh-at-emory.edu
Website: http://electronmicroscopy.emory.edu

Applications will be reviewed as they are received and until the
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Dear Listers,

Greeting from the Netherlands

I am looking for a Tecnai F20 and/or a Phillips CM20 specimen holder
to perform some basic mechanical tests (e.g., stiffness measurement,
heat expansion coefficient determination etc.). The holder does not
need to be in working condition, nor does it need to be new. It must,
however, be mechanically sound (that is, not physically damaged).

If any of you know how I could procure such holder (e.g., I could buy
it from someone that wants to get rid of a broken one) please let me
know.

Many thanks for your help.

Kind regards,

Arturo Tejada

--

===================================================
Dr. Arturo Tejada
Assistant Professor
Mechatronic System Design Group
Precision and Microsystems Engineering Dpt.
Mekelweg 2, Room G-1-300
2628 CD, Delft, The Netherlands
Phone: +31-15-27-81550
Fax: +31-15-27 89475
www.tejadaruiz.net
===================================================

==============================Original Headers==============================
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9, 29 -- Subject: [TEM] Looking for a specimen holder
9, 29 -- From: Arturo Tejada {a.tejadaruiz-at-tudelft.nl}
9, 29 -- To: Microscopy-at-microscopy.com
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Email: svmcphie-at-utmb.edu
Name: svetla stoilova-mcphie

Organization: University fo texas Medical Branch

Title-Subject: [Filtered] Postdoctoral position in Cryo-EM

Message: A postdoctoral position is available in my group
www.svetla-mcphie-cryoem.com to study the structure of membrane-bound
coagulation proteins by Cryo-electron microscopy. This position is part
of a collaboration with Professor B.M. Pettitt utilizing dynamic
simulation refinements combined with direct structure determination by
Cryo-EM.
The ideal candidate should be highly motivated and have a PhD degree in
the physical or biological sciences, with backround in macromolecular
structure determination. Experience with unix computing systems and
programming will be an advantage. Candidates who would like to learn
Cryo-EM, structure analysis and molecular modeling are also encouraged
to apply.
The University of Texas Medical Branch at Galveston and the Sealy Center
for Structural Biology and Molecular Biophysics are fully equipped for
state of the art structure determination of macromolecular complexes by
Cryo-EM: http://www.scsb.utmb.edu/facilities/cryo/ UTMB Interested
candidates should send a CV including publications list, and names and
contacts for 3 references to svmcphie-at-utmb.edu.
The position is offered initially for two years. UTMB is an equal
opportunity employer and the postdoctoral salary will be according to
NIH standard scales for postdoctoral fellows:
http://grants2.nih.gov/grants/guide/notice-files/NOT-OD-10-047.html
commensurate with experience and UTMB regulations.

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Hello,

We are in the process of measuring particles (actually metal nodule type objects that are seen from the top-down) and we are looking for information such as: how many are in the image, how much area they take up (each and in total), and average size. Our current solution for this is with ImageJ, since it is free and simple to use for this sort of thing. The issue we have with it is the time it takes for each image to be processed. Does anyone know of any other sort of automated software to handle this sort of thing?

We've tried the thresholding function to separate the nodules, but the grayscale of the image and the closeness of the objects don't allow us to get accurate separation of them for ImageJ to auto-magically measure them. So, our solution has been to free hand select each one and track/measure with the ROI manager.

I'm hoping that some of you have had this issue before and maybe some ideas to get through it. To give an idea of the time saving need, we will be processing at least two of these types of images a day for the next couple years (at least it's steady work huh?) and at the rate we're going now, it takes about 45min to an hour per image.

Thanks in advance,

Andrew Ornelas
Failure Analysis Lab Technician

Intertek AIM | intertek.com/aptech

Valued Quality. Delivered.
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12, 35 -- From andrew.ornelas-at-intertek.com Thu Sep 27 13:42:49 2012
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12, 35 -- From: "Andrew Ornelas Intertek" {andrew.ornelas-at-intertek.com}
12, 35 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
12, 35 -- Subject: SEM Image Analysis for Particle Size and Distribution
12, 35 -- Thread-Topic: SEM Image Analysis for Particle Size and Distribution
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Andrew,

It seems to me that "Watershed Segmentation" function in FoveaPro add-on
to the Photoshop should do just the trick that you need for separating
overlapping grains, but I am not sure if FoveaPro has any macro
programming capabilities to completely automate your measurements.

Take a look here: http://reindeergraphics.com/science.html - scroll down
to the second example.

It is extremely likely however, that someone already written a plugin
for ImageJ with functionality equivalent to "Watershed Segmentation" -
search here: http://rsbweb.nih.gov/ij/plugins/index.html If such plugin
exists, then you should be able to use macro-programming of ImageJ to
get fully automated processing...

Good luck!

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
Web: www.partbeamsystech.com
Web: www.freudlabs.com

On 9/27/2012 2:43 PM, andrew.ornelas-at-intertek.com wrote:
} ----------------------------------------------------------------------------
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} Hello,
}
} We are in the process of measuring particles (actually metal nodule type objects that are seen from the top-down) and we are looking for information such as: how many are in the image, how much area they take up (each and in total), and average size. Our current solution for this is with ImageJ, since it is free and simple to use for this sort of thing. The issue we have with it is the time it takes for each image to be processed. Does anyone know of any other sort of automated software to handle this sort of thing?
}
} We've tried the thresholding function to separate the nodules, but the grayscale of the image and the closeness of the objects don't allow us to get accurate separation of them for ImageJ to auto-magically measure them. So, our solution has been to free hand select each one and track/measure with the ROI manager.
}
} I'm hoping that some of you have had this issue before and maybe some ideas to get through it. To give an idea of the time saving need, we will be processing at least two of these types of images a day for the next couple years (at least it's steady work huh?) and at the rate we're going now, it takes about 45min to an hour per image.
}
} Thanks in advance,
}
} Andrew Ornelas
} Failure Analysis Lab Technician
}
} Intertek AIM | intertek.com/aptech
}
}
} Valued Quality. Delivered.
} ------------------------------------------------------------------------------
} CONFIDENTIALITY NOTICE
}
} This email may contain confidential or privileged information, if you are not the intended recipient, or the person responsible for delivering the message to the intended recipient then please notify us by return email immediately. Should you have received this email in error then you should not copy this for any purpose nor disclose its contents to any other person.
}
} http://www.intertek.com
}
}
} ==============================Original Headers==============================
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} 12, 35 -- To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
} 12, 35 -- Subject: SEM Image Analysis for Particle Size and Distribution
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Dear Listservers,

It is with great sadness that I write regarding the death of a
long-time colleague and friend, Chris Cathcart.

About a year ago, Chris was diagnosed with a brain tumor. However,
after successful surgery, he went back to work and picked up his
normal life. Apparently, the tumor re-emerged, aggressively, about a
month ago, and Chris lost the battle last Thursday night.

He will long be remembered for his energy, his humor, his knowledge,
and his enthusiasm.

Condolences can be sent to his sister:
Pamela LeBlanc
7 Kenins Crescent
Kanata, Ontario
K2S-2S7
Canada.
pleblanc-at-johnsheainsurance.com

Best regards,
Barbara Foster, President and Sr. Consultant

Microscopy/Microscopy Education
P: (972)924-5310
W: www.MicroscopyEducation.com

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Hi Listers,

I've realized how popular of an issue this is after the many replies. Thank you all for your input, and I am still going through each email for their valuable information. Some have asked to see some example images so I've put some up on my google drive: https://docs.google.com/folder/d/0Bz_h0BMsA6KSTUplSDRXVVEyQTg/edit

Hopefully this helps illustrate what my email was trying to explain. If you try to view the image, please download the image first because the "google viewer" seems to make it one big blur.

Thank you again for everyone's help and I hope that we can use this info in the future for any upcoming lister's predicaments.

Thanks,

Andrew Ornelas
Failure Analysis Lab Technician

Intertek AIM | intertek.com/aptech

Valued Quality. Delivered.
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Email: r-bleher-at-northwestern.edu
Name: Reiner Bleher

Organization: Northwestern University

Title-Subject: [Filtered] Research Position Available

Message: Dear Colleagues,

A Cryo-EM Research Position is available at NUANCE Center

Northwesterns NUANCE Center (Northwestern University's Atomic and Nanoscale Characterization
Experimental Center, http://www.nuance.northwestern.edu) has an immediate opening for a postdoctoral
scholar or senior research associate with a background in cryo-bio S/TEM imaging/analysis of
assembly/organization of soft and hybrid nanostructures, such as patterned DNA/lipids,
nanoparticle-DNA/protein assemblies, and related techniques.
The position is part of extensive cross-disciplinary and collaborative materials research activities
at Northwestern. Thus, the candidate has ample opportunities to learn and contribute to
analytical-/in-situ-, and cryo-EM in materials science, and bio- and nano-technologies. The NUANCE
Center and sister facilities are well equipped with modern S/TEMs, SEMs and extensive specimen
preparation capabilities, ranging from focused ion beam (FIB) to cryo-transfer with LN2 stages. The
position requires a PhD in biological/physical sciences/engineering. Respective experience in
cryo-EM techniques and computation/simulations is essential. The position is available immediately
for at least two years, with the possibility for mutual extension. Salary and compensation would
commensurate with experience, up to: $ 55,000-$65,000 per year. A select candidate is eligible for
an IIN Postdoctoral Fellowship (International Institute for Nanotechnology), which offers additional
significant compensation on top of the core stipend.

To apply, please send your CV and a list of three references with email addresses and telephone
numbers to Reiner Bleher, r-bleher-at-northwestern.edu.

Sincerely,

Dr. Reiner Bleher
Research Assistant Professor
NUANCE-Center / EPIC
Department of Materials Science and Engineering
Northwestern University

http://www.nuance.northwestern.edu
http://www.nuance.northwestern.edu/epic/index.html

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Title-Subject: [Filtered] magnetic nano particles

Message: I have a user who wants to view "magnetic nano particles" on our TEM....I do not feel
comfortable putting anything magnetic by/ in scope - thoughts???
thanks
sue

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Email: z.zhou-at-lboro.ac.uk
Name: Zhaoxia

Organization: Loughborough University

Title-Subject: [Filtered] Electron diffraction vs X-ray diffraction

Message: Dear Listers,

Can I have your opinions on the following questions---

What are the advantages/disadvantages of Electron diffraction compared to X-ray diffraction in terms
of identifying crystal structures in material sciences? How accurate are both techniques now days?

For electron diffraction, 2000FX and TecnaiF 20 are the TEMs we use. For X-ray diffraction, the
Bruker D8.

Zhou

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Email: tkremer-at-ipstesting.com
Name: Tom Kremer

Organization: Integrated Paper Services

Title-Subject: [Filtered] Oxford INCA

Message: Folks,
I recently acquired an Oxford INCA system for our SEM. For anyone with this system, I have a problem
needing some advice. This system is operating without "Autocolumn"; therefore I manually enter
magnification, voltage and working distance in a separate window called "Microscope Control" prior
to acquiring images and spectra. However, if I forget to modify the settings at a new sampling
location, which is too easy to do, the wrong values are recorded with the image or spectra. The
tutorial says "No problem", just enter the correct values in the assigned fields and click on
"Apply". So here's the problem: the graphic of the screen in the tutorial shows fields that can be
modified after Magnification and Accelerating Voltage with the "Apply" button below them. However,
my system screen simply has a field with Magnification and Accelerating Voltage that CANNOT be
modified. There is a button, actually two buttons in a field beneath called "Restore", one for
Column and one for Stage. Clicking on the Column button will assign the mag and voltage values
associated with the current image to the values in the Microscope Control Window. That is backward
from what I need to do. Your help will be appreciated.

Hopefully this problem will go away if my service engineer can get the program to talk to the SEM
correctly.

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Yes, you should feel uncomfortable! After all, all the lenses are magnetic.

I require users to use membrane coated, double grids - put the particles in the middle of the sandwich. It is safe that way because the particles would not be able to escape.

Zhaojie Zhang
Director, Jenkins Microscopy Facility
University of Wyoming
Laramie, WY 82071
________________________________________
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Name: Susan Van Horn

Title-Subject: [Filtered] magnetic nano particles

Message: I have a user who wants to view "magnetic nano particles" on our TEM....I do not feel
comfortable putting anything magnetic by/ in scope - thoughts???
thanks
sue

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Email: Zaluzec-at-Microscopy.Com

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On Sep 28, 2012, at 2:48 PM, microscopylistserver-
noreply-at-microscopy.com wrote:

} Email: z.zhou-at-lboro.ac.uk
} Name: Zhaoxia
}
} Organization: Loughborough University
}
} Title-Subject: [Filtered] Electron diffraction vs X-ray diffraction
}
} Message: Dear Listers,
}
} Can I have your opinions on the following questions---
}
} What are the advantages/disadvantages of Electron diffraction
} compared to X-ray diffraction in terms
} of identifying crystal structures in material sciences? How accurate
} are both techniques now days?
}
} For electron diffraction, 2000FX and TecnaiF 20 are the TEMs we use.
} For X-ray diffraction, the
} Bruker D8.
}
} Zhou

Dear Zhou,
I cannot speak for XRD, but one difference between the two is the
stronger interaction of electrons with the specimen. This can be
either an advantage, when there are only very small crystals, or a
disadvantage, when dynamical effects from multiple scattering perturb
the intensities. Convergent beam electron diffraction (CBED) can
provide a great deal of information, but it requires a pretty large
dose--not often a problem in materials science, but something to be
aware of. There are many good references to both selected area ED and
CBED, and you should find out about accuracy from one of them.
Yours,
Bill

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Dear Sue,

I can offer a couple of insights that might help.

I've observed ferromagnetic nano-particle work in several TEMs based on
similar electron optical column design, namely Philips/FEI CMs (CM30 and
CM300F) and Tecnai (F20). We have not observed any significant problems
with these instruments because the magnetic field distribution within the
objective is relatively symmetric, i.e. the magnetic field gradient at the
specimen level is very small/zero. A magnetic particle will feel a torque,
but no overall force, if the B-field is constant in space. A nano-particle
will either mechanically twist or, if the magnetisation is relatively easy
to change (low coercivity), the magnetisation will align to the field of
the lens, i.e. paramagnetic behaviour.

Magnetic nano-particles placed in highly asymmetric field configurations
are a problem because the field gradient is significant and the particles
will move towards the direction in which the field gradient is largest,
i.e. towards the lens. It was not uncommon to find a nice little collection
of magnetic particles sitting on the pole-piece of VG STEMs for example.
These instruments have a single pole-piece and a highly asymmetric field as
a consequence.

If your microscope has a relatively symmetric field distribution, you
should have no problems, e.g. FEI TWIN-lens designs. I have no experience
of JEOL and LEO/Zeiss instruments, so cannot comment. If you're unsure, ask
your EM manufacturer.

If you're still not convinced, then say "no" by all means. Caution is the
better part of valour as we say.

Yours, Jon

} X-from: susan.vanhorn-at-stonybrook.edu ()
}
} This Question/Comment was submitted to the Microscopy Listserver using
} the WWW based Form at http://www.microscopy.com/MLFormMail.html
}
} Email: susan.vanhorn-at-stonybrook.edu
} Name: Susan Van Horn
}
} Title-Subject: [Filtered] magnetic nano particles
}
} Message: I have a user who wants to view "magnetic nano particles" on our
} TEM....I do not feel comfortable putting anything magnetic by/ in scope -
} thoughts??? thanks sue
}
} ---------------------------------------------------------------------------

==============================Original Headers==============================
9, 28 -- From jsb43-at-hermes.cam.ac.uk Sun Sep 30 08:44:14 2012
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I my year and a half at Western Digital Media, where all the sample are fine grained magnetic recording materials. I was told that the SEMS needed the pole pieces cleaned every several years. The TEM never had that problem, but the TEM was only two years old when I left. The SEMs see a much larger number of magnetic grains.

John Mardinly, ASU
}
}
} ----------------------------------------------------------------------------
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}
} Dear Sue,
}
} I can offer a couple of insights that might help.
}
} I've observed ferromagnetic nano-particle work in several TEMs based on
} similar electron optical column design, namely Philips/FEI CMs (CM30 and
} CM300F) and Tecnai (F20). We have not observed any significant problems
} with these instruments because the magnetic field distribution within the
} objective is relatively symmetric, i.e. the magnetic field gradient at the
} specimen level is very small/zero. A magnetic particle will feel a torque,
} but no overall force, if the B-field is constant in space. A nano-particle
} will either mechanically twist or, if the magnetisation is relatively easy
} to change (low coercivity), the magnetisation will align to the field of
} the lens, i.e. paramagnetic behaviour.
}
} Magnetic nano-particles placed in highly asymmetric field configurations
} are a problem because the field gradient is significant and the particles
} will move towards the direction in which the field gradient is largest,
} i.e. towards the lens. It was not uncommon to find a nice little collection
} of magnetic particles sitting on the pole-piece of VG STEMs for example.
} These instruments have a single pole-piece and a highly asymmetric field as
} a consequence.
}
} If your microscope has a relatively symmetric field distribution, you
} should have no problems, e.g. FEI TWIN-lens designs. I have no experience
} of JEOL and LEO/Zeiss instruments, so cannot comment. If you're unsure, ask
} your EM manufacturer.
}
} If you're still not convinced, then say "no" by all means. Caution is the
} better part of valour as we say.
}
} Yours, Jon
}
} } X-from: susan.vanhorn-at-stonybrook.edu ()
} }
} } This Question/Comment was submitted to the Microscopy Listserver using
} } the WWW based Form at http://www.microscopy.com/MLFormMail.html
} }
} } Email: susan.vanhorn-at-stonybrook.edu
} } Name: Susan Van Horn
} }
} } Title-Subject: [Filtered] magnetic nano particles
} }
} } Message: I have a user who wants to view "magnetic nano particles" on our
} } TEM....I do not feel comfortable putting anything magnetic by/ in scope -
} } thoughts??? thanks sue
} }
} } ---------------------------------------------------------------------------
}
}
} ==============================Original Headers==============================
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Name: bernd.zechmann-at-uni-graz.at

Organization: University of Graz

Title-Subject: [Filtered] Normalization of immunogoldlabeling

Message: Dear members of the list server,

During the last review process of a manuscript showing quantitative immunogold data collected by TEM
the question arose as to why we are not normalizing the gold particle density of the desired antigen
to an internal control protein such as actin through double labeling. Apparently this is especially
crucial when using different samples cultivated under different conditions. Looking through the
literature in my field (plant sciences) I was not able to find publications where quantitative
immunogold labeling data was normalized against an internal standard. So my questions are:
1.) Why is this procedure, which is commonly used during rt-PCR and microarrays, not used for
quantitative immungold labeling?
2.) Is anybody aware of publications where gold particle density was normalized against an internal
standard e.g. actin?
3.) If such a normalization were to be performed which protein could serve as a control protein in
plants, in my opinion it would need to occur in all cell compartments?

In my case I've analysed immunogold labeling density after the incubation of ultrathin sections with
primary (against the desired antigen) and secondary gold conjugated antibodies in different cell
compartments. Then I've compared labeling density in the different cell compartments between
different control and treated/stressed samples.

Thanks for your help in advance.

Bernd

++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++

Dr. Bernd Zechmann
Institute of Plant Sciences
University of Graz
Schubertstra�e 51
8010 Graz
Austria/Europe
Tel.: ++43/316/380/5635
Fax.: ++43/316/380/9880

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Hi Bernd,
�
1) As I understand it the quantification of actin is used to normalize the amount of material analyzed. So if you quantify a protein you express it as �g of this protein per mg of total material (cell extract, cell number, whatever).
In your case you normalize the density of labeling�by expressing your results per cm� for example.
�
2) I think it is dangerous to compare the density of 2 labelings together. You can use double labelings to compare the localization of 2 epitopes but not for quantification. Each labeling has its own characteristics and one condition may show an increase of labeling not because there are more epitopes per cm� but because (f.ex.) the epitope is better presented, or has been relocalized in a comparment where the antibodies have better access to it, or because it was present in a dense complex and after treatment it is not associated with the complex anymore,�or the protein has refolded.�
So after treatment you see that your labeling for actin is more dense (gold particles per cm�), can you conclude anything about another labeling? What is the relationship with the other epitope? There is simply none. It simply doesn't help to quantify actin at the same time as your epitope of interest because you cannot conclude for one from the results of the other.
�
Aknowledgement: I am a biologist and I do labelings in LM but not in EM. I am just using my common sense here, so it would still be interesting to hear from those-who-know.
�
Macht's gut
Stephane
�
�
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Email: bernd.zechmann-at-uni-graz.at
Name: bernd.zechmann-at-uni-graz.at

Organization: University of Graz

Title-Subject: [Filtered] Normalization of immunogoldlabeling

Message: Dear members of the list server,

During the last review process of a manuscript showing quantitative immunogold data collected by TEM
the question arose as to why we are not normalizing the gold particle density of the desired antigen
to an internal control protein such as actin through double labeling. Apparently this is especially
crucial when using different samples cultivated under different conditions. Looking through the
literature in my field (plant sciences) I was not able to find publications where quantitative
immunogold labeling data was normalized against an internal standard. So my questions are:
1.) Why is this procedure, which is commonly used during rt-PCR and microarrays, not used for
quantitative immungold labeling?
2.) Is anybody aware of publications where gold particle density was normalized against an internal
standard e.g. actin?
3.) If such a normalization were to be performed which protein could serve as a control protein in
plants, in my opinion it would need to occur in all cell compartments?

In my case I've analysed immunogold labeling density after the incubation of ultrathin sections with
primary (against the desired antigen) and secondary gold conjugated antibodies in different cell
compartments. Then I've compared labeling density in the different cell compartments between
different control and treated/stressed samples.

Thanks for your help in advance.

Bernd

++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++

Dr. Bernd Zechmann
Institute of Plant Sciences
University of Graz
Schubertstra�e 51
8010 Graz
Austria/Europe
Tel.: ++43/316/380/5635
Fax.: ++43/316/380/9880

� Login Host: 143.50.114.113
� Listserver Email Form V - 20120416
---------------------------------------------------------------------------

--
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===========================================
Dr. Nestor J. Zaluzec
797 Bonnie Brae Ct.
Bolingbrook, Illinois 60440, USA
Tel:1-630-739-1160
Alternate: 1-530-NES-TORZ (1-530-637-8679) has Voice Mail
Email: Zaluzec-at-Microscopy.Com

iChat:Zaluzec-at-AIM
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===========================================

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Hi,
As usual with this things, potential
answers depend on the research question.

Often, immunogold is used to identify an
structure (what is this funny looking blob?) or
localize an antigen (is my protein is present in
the lysosome?). For these purposes, it should be
fine to count label densities over appropriate
regions, with the usual no primary control.

It is more complex if you want to use
immunogold to measure the amount of antigen in
some region. It has always been my impression
that, in sections, the relation between antigen
amount and antibody binding is non-linear (or
linear over a quite restricted range). This
limits the precision fairly extensively.
Therefore, I think no amount of normalization is
going to magic away this problem and give you
perfect quantification. I'd say the best you can
get is a reproducible approximation.

If you want to push the PCR analogy,
most careful qPCR runs use several genes to
normalize, not just one gene. Following this, you
would need to say mix three control primary abs,
detect them all with one size gold particle
secondary, and then detect yours with another.
While this sounds cute in principle, in practice
I don't think it work well enough to justify the
effort (or example steric hinderance is likely to
be a big problem).

One should be able to control for the
different conditions by reporting ratios of
labeling, rather than the absolute amounts. So
the density of gold particles in mitochondria to
chloroplasts say would be the parameter reported.
This seems robust to most sorts of sample prep
vagaries (quality of fixation, so on).

Hope this helps,
Tobias

}
}
} X-from: bernd.zechmann-at-uni-graz.at ()
}
} This Question/Comment was submitted to the Microscopy Listserver
} using the WWW based Form at http://www.microscopy.com/MLFormMail.html
} ---------------------------------------------------------------------------
} Remember this posting is most likely not from a Subscriber, so when replying
} please copy both bernd.zechmann-at-uni-graz.at as
} well as the Microscopy Listserver
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}
} Email: bernd.zechmann-at-uni-graz.at
} Name: bernd.zechmann-at-uni-graz.at
}
} Organization: University of Graz
}
} Title-Subject: [Filtered] Normalization of immunogoldlabeling
}
} Message: Dear members of the list server,
}
} During the last review process of a manuscript
} showing quantitative immunogold data collected
} by TEM
} the question arose as to why we are not
} normalizing the gold particle density of the
} desired antigen
} to an internal control protein such as actin
} through double labeling. Apparently this is
} especially
} crucial when using different samples cultivated
} under different conditions. Looking through the
} literature in my field (plant sciences) I was
} not able to find publications where quantitative
} immunogold labeling data was normalized against
} an internal standard. So my questions are:
} 1.) Why is this procedure, which is commonly
} used during rt-PCR and microarrays, not used for
} quantitative immungold labeling?
} 2.) Is anybody aware of publications where gold
} particle density was normalized against an
} internal
} standard e.g. actin?
} 3.) If such a normalization were to be performed
} which protein could serve as a control protein in
} plants, in my opinion it would need to occur in all cell compartments?
}
} In my case I've analysed immunogold labeling
} density after the incubation of ultrathin
} sections with
} primary (against the desired antigen) and
} secondary gold conjugated antibodies in
} different cell
} compartments. Then I've compared labeling
} density in the different cell compartments
} between
} different control and treated/stressed samples.
}
} Thanks for your help in advance.
}
} Bernd
}
} ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
}
} Dr. Bernd Zechmann
} Institute of Plant Sciences
} University of Graz
} Schubertstra�e 51
} 8010 Graz
} Austria/Europe
} Tel.: ++43/316/380/5635
} Fax.: ++43/316/380/9880
}
}
} Login Host: 143.50.114.113
} Listserver Email Form V - 20120416
} ---------------------------------------------------------------------------
}
}
}
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} ---===[|]===---
}
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} 797 Bonnie Brae Ct.
} Bolingbrook, Illinois 60440, USA
} Tel:1-630-739-1160
} Alternate: 1-530-NES-TORZ (1-530-637-8679) has Voice Mail
} Email: Zaluzec-at-Microscopy.Com
}
} iChat:Zaluzec-at-AIM
} Skype:Zaluzec
} ===========================================
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}
}
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} Do not reply to this message it is from
} the Microscopy Listserver NO-REPLY forwarding
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Hello All,

The Biological Imaging Facility (BIF) at Northwestern University has an opening for a Microscopy & Imaging Specialist at the Research Staff/Research Professor level. BIF serves as an intellectual and experimental resource for biological imaging on the Evanston campus, with over 400 users of photonic and electron microscopy, specimen preparation, and printed output.

The educational requirement for this position is a graduate degree in cell biology, chemistry, biophysics or related field. Preference will be given to applicants with extensive research experience in photonic and/or electron microscopy, advanced live cell imaging techniques, biological specimen and tissue preparation, as well as digital image analyses. The candidate should be familiar with maintenance and calibration of microscopes. Experience with grant writing is highly desirable.

The candidate must have the ability to work effectively with a wide diversity of people and the ability to work both independently and cooperatively. Strong analytical and organizational skills coupled with excellent interpersonal and communication skills (both oral and written) are essential. Also highly desirable are good manual dexterity, and good technical aptitude. The Specialist will also be involved in teaching in various microscopy workshops organized by BIF.

Salary commensurate with skills and experience.

Major responsibilities of this position include acting as an expert resource to faculty, staff, and students for experimental design, troubleshooting specialized microscopy applications, and image quantitation. The Specialist will be expected to assist the operations director with grant preparation. Additional responsibilities include: routine instrument maintenance, new user training, and assistance with other aspects of facility operation as necessary. Position responsibilities may also include interaction with work-study students, and other duties as assigned.

Applicants should complete an online application at http://www.northwestern.edu/hr/careers/index.html (Microscopy & Imaging Specialist - Job ID 19824).

Applications will be accepted and reviewed until position is filled.

William A. Russin Ph. D.
Director, Biological Imaging Facility
Department of Neurobiology
Northwestern University
2205 Tech Drive
Evanston, IL 60208
w-russin-at-northwestern.edu

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Dear Listers,
�
I have experience in TEM and SEM but none in EPMA.
I would like to know what kind of gun is used: W, LaB6, FEG?
I would like to know what X-Y resolution can be attained.
I suppose that it is not possible to go down too much with the kV otherwise you cannot measure lighter elements so one has probably to�take strongly in consideration the interaction volume inside the specimen (Z resolution), which increases with the kV.
�
To take an example, what X-Y (and Z) resolution can be expected using a Cameca SX 100 instrument�with an aluminosilicate specimen ("light" mineral).
Does it make sense to expect to be able to resolve different minerals in particles 3-10�m in size?
�
Any consideration welcome.
Many thanks in advance.
�
Stephane

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Email: lamiller-at-illinois.edu
Name: Lou Ann Miller

Organization: Frederick Seitz Materials Research Laboratory

Title-Subject: [Filtered] Full Day workshop at MRL

Message: Greetings,

We at the Materials Research Laboratory at the U of Illinois, are putting on a day of tutorials on
basic biological structure, and introduction into both optical and electron techniques and morphology.

Time: 8am - 5 pm

Activities: Tutorials, Vendor Fair, Tours of MRL, Open wet lab for parasitology, Breakfast Break,
Lunch and afternoon break.

Where: 190 ESB, at the corner of Springfield and Goodwin, Urbana Illinois.(Map on web page)

Cost: Before October 15th, $25/ participant and includes all food. After Oct 15 it is $35

Speakers:

Basic & Comparative Histology: Dr. Kuldeep Singh & Dr. Adam Stern, College of Veterinary Medicine
Parasitology: Dr. Allen Paul, College of Veterinary Medicine
Entomology: Dr. Jim Nardi, Department of Entomology
Comparing Light and Electron Microscopy: Dr. Rex Hess, Colleges of Medicine & Veterinary Medicine
Biological Imaging Techniques and Electron Microscopy Images: Lou Ann Miller, MRL/Engineering

Where to register, get information, the schedule and the downloadable flyer:
http://mrl.illinois.edu/news/conferences-workshops/biological-structures-electron-optical-imaging-workshop

Vendors:
We still have some room at the vendor fair. Currently we have several instruments that will be
demonstrated, you can contact me (Lou Ann) or visit the vendor Registration page at:
https://my.mrl.illinois.edu/eventreg/register.asp

** Many projects involve both material and biological substances, this is a chance for material
scientists to get an introduction into basic histology and electron microscopy tissue structures.
This workshop is also suitable for biologists as an introduction into electron microscopy.
Undergrads, grad students, post docs, faculty and those just plain curious, are all welcome to attend.

{ { { { { { { { { { { { { { { { { { {} } } } } } } } } } } } } } } } } }
Lou Ann Miller
Biological Electron Microscopy
Frederick Seitz Materials Research Laboratory
Room 125
104 South Goodwin Ave
Urbana, IL 61801
Campus mail code: MC-230

217-244-1567
lamiller-at-illinois.edu
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Dear Stephane,
Roughly speaking, I'd say that SEM and EPMA are the same type of instrument today, but SEM is optimized for resolution, and EPMA is optimized for needs of X-ray microanalysis (i.e. stability, specimen positioning, ports for a number of WDS spectrometers). Any kind of a gun can be used, with exception of cold FEG (not stable enough). As for spatial analytical resolution, it is exactly the same as for SEM (same physics.) Energy (wavelength) resolution of WDS is, of course, much better than that of EDS. As for accelerating voltage, on EPMA with Schottky gun you can go as low as you wish (the same as for SEM/EDS). For aluminosilicates you can use 3-4 kV; the interaction volume I believe (but I am not sure) will be about 100-200 nm. Lighter elements are good for high resolution analytical work. As for particles 3-10�m in size - they are huge; no problems at all with qualitative analysis.

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: Wednesday, October 03, 2012 1:31 AM
} To: Dusevich, Vladimir
} Subject: [Microscopy] Question about Electron Probe Micro-Analysis
}
}
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} Dear Listers,
}
} I have experience in TEM and SEM but none in EPMA.
} I would like to know what kind of gun is used: W, LaB6, FEG?
} I would like to know what X-Y resolution can be attained.
} I suppose that it is not possible to go down too much with the kV otherwise
} you cannot measure lighter elements so one has probably to�take strongly in
} consideration the interaction volume inside the specimen (Z resolution),
} which increases with the kV.
}
} To take an example, what X-Y (and Z) resolution can be expected using a
} Cameca SX 100 instrument�with an aluminosilicate specimen ("light" mineral).
} Does it make sense to expect to be able to resolve different minerals in
} particles 3-10�m in size?
}
} Any consideration welcome.
} Many thanks in advance.
}
} Stephane
}
}
} ==============================Original
} Headers==============================
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4, 29 -- From DusevichV-at-umkc.edu Wed Oct 3 09:29:15 2012
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4, 29 -- Subject: RE: [Microscopy] Question about Electron Probe Micro-Analysis
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Stephane

Let me add a couple of comments to the information Vladimir gave.

The largest difference between SEM and EPMA is not the electron gun and
column, it is the issue of detectors... EPMA utilizes wavelength
dispersive spectrometers with gas detectors whereas SEM utilizes a solid
state energy dispersive spectrometer+detector. Of course many EPMAs also
have an EDS detector, and a few SEMs have an add-on WDS.

You mentioned specifically measurement of light element contents.

1. The spectral resolution of WDS is on the order of 10 times or more
better than EDS, so that interferences by L and M lines of elements
present in the sample falling on or near the light element of interest
can in many/most cases will not cause errors, whereas with EDS this may
not be the case.
2. EPMA-WDS traditionally utilizes standards, so that quality control is
possible (is the analytical total close to 100 wt%?), whereas SEM-EDS
traditionally does not utilize explicit standards and normalizes the
results to 100 wt%, so quality control can be an issue.

John Fournelle
UW Electron Microprobe Lab
Madison, WI 53706

On 10/3/12 9:33 AM, DusevichV-at-umkc.edu wrote:
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} Dear Stephane,
} Roughly speaking, I'd say that SEM and EPMA are the same type of instrument today, but SEM is optimized for resolution, and EPMA is optimized for needs of X-ray microanalysis (i.e. stability, specimen positioning, ports for a number of WDS spectrometers). Any kind of a gun can be used, with exception of cold FEG (not stable enough). As for spatial analytical resolution, it is exactly the same as for SEM (same physics.) Energy (wavelength) resolution of WDS is, of course, much better than that of EDS. As for accelerating voltage, on EPMA with Schottky gun you can go as low as you wish (the same as for SEM/EDS). For aluminosilicates you can use 3-4 kV; the interaction volume I believe (but I am not sure) will be about 100-200 nm. Lighter elements are good for high resolution analytical work. As for particles 3-10�m in size - they are huge; no problems at all with qualitative analysis.
}
} Vladimir M. Dusevich, Ph.D.
} Electron Microscope Lab Director
} 371 School of Dentistry
} 650 E. 25th Street
} Kansas City, MO 64108-2784
}
} } -----Original Message-----
} } From: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} } Sent: Wednesday, October 03, 2012 1:31 AM
} } To: Dusevich, Vladimir
} } Subject: [Microscopy] Question about Electron Probe Micro-Analysis
} }
} }
} }
} }
} } ----------------------------------------------------------------------------
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} }
} } Dear Listers,
} }
} } I have experience in TEM and SEM but none in EPMA.
} } I would like to know what kind of gun is used: W, LaB6, FEG?
} } I would like to know what X-Y resolution can be attained.
} } I suppose that it is not possible to go down too much with the kV otherwise
} } you cannot measure lighter elements so one has probably to take strongly in
} } consideration the interaction volume inside the specimen (Z resolution),
} } which increases with the kV.
} }
} } To take an example, what X-Y (and Z) resolution can be expected using a
} } Cameca SX 100 instrument with an aluminosilicate specimen ("light" mineral).
} } Does it make sense to expect to be able to resolve different minerals in
} } particles 3-10�m in size?
} }
} } Any consideration welcome.
} } Many thanks in advance.
} }
} } Stephane
} }
} }
} } ==============================Original
} } Headers==============================
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} 4, 29 -- From: "Dusevich, Vladimir" {DusevichV-at-umkc.edu}
} 4, 29 -- To: "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
} 4, 29 -- Subject: RE: [Microscopy] Question about Electron Probe Micro-Analysis
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--
John Fournelle
Senior Scientist
Director, Electron Probe and SEM Lab
University of Wisconsin Dept of Geoscience
Madison, Wisconsin 53706
Office 608-262-7964 Cell 608-438-7480

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Dear all,

It looks that one essential advantage of EPMA (or WDS) on EDS is still
missing.

On the hand the EDS detector receive all the X-ray photons within the
collection angle from all elements present under the electron probe. On
the other hand the counting rate si limited be the
processor/multichannel analyzer. When analyzing samples with major and
minor elements - let say a steel or high temperature alloys for instance
- the major element (Fe, Ni...) reach a number of counts that give an
excellent statistical relevancy. At contrary minor elements like C or
addition elements (Mo, W, Ta, Zr...) have poor counts even after a long
acquisition and their quantification remains quite uncertain.

In an EPMA each detector sees only photons of one element at a time.
Thus an educated operator will choose a weak line (or second order line)
for the major element to bring its count rate down to the level of that
one of the most intense line of the minor elements. Then he will boost
the probe current to get a high count rate for everybody. This
dramatically improves the statistical relevancy and composition accuracy
for light elements. It also brings better detectability of traces
elements towards tens of ppm or even ppm instead of the classical 0.1%
of EDS.

Philippe Buffat
EPF-Lausanne Switzerland
AGH-Krakow Poland

==============================Original Headers==============================
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***************************************************************************************
Forwarded from "Ask a Microscopist"
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**any reply should go directly to the poster**
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Using the "reply" function in your email does *not* send your answer
to the person asking the question.
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****************************************************************************************
} realname - Jagdeep Kaur
} Email - jkaur-at-danforthcenter.org
} ORGANIZATION - Donald Danforth Plant Science Center
} EDUCATION - Graduate College
} LOCATION - St. Louis, Missouri, USA
} SUBJECT_OF_QUESTION - Help with confocal
} microscopy to image Unitex 2B stained leaves
} QUESTION - Hi All-
}
} I wish to image my transgenic wheat leaves
} infected with rust fungi (leaf and stripe rust
} fungus-Puccinia sps.) using a recent protocol
} described by Sorensen et al. (2012) in Journal
} Mycologia (in press). Basically leaves are
} stained with Unitex2B dye (after clearing
} chlorophyll and other reagents) and need to
} imaged on Zeiss LSM 510 confocal microscope
} equipped with with a Plan-Apochromat 63�~/1.4
} oil immersion objective. Excitation of
} Uvitex-2B stained structures was done with a Mai
} tai two-photon laser at wavelengths of 720 nm.
} Emitted light was scanned with filters settings
} for short passing of 685 nm.
} Unfortunately our confocal microscope does not
} have this filter so I am looking for other
} facilities who could do this for me???
} Thanks in advance for help and time.
} Jagdeep

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X-from: George.Theodossiou-at-st-annes.ox.ac.uk ()

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Email: George.Theodossiou-at-st-annes.ox.ac.uk
Name: George Theodossiou

Organization: University of Oxford

Title-Subject: [Filtered] Digital Micrograph - Scripted Serial Acquisitions for EELS and Spectrum
Imaging

Message: Hi Listers,

I've been a long time 'listener' to this forum and enjoy reading the posts. Now its my turn to ask
all of you for information and assistance.

I'm after some scripting help with Gatan's Digital Micrograph.
I'm using a VG HB501 STEM with a Gatan Model 666 spectrometer and a Pixis camera to record the EELS
spectrum. The microscope also has a Digiscan to control the beam
Works a treat....however.....I am observing time dependent processes and would like sequential EELS
spectra and Spectrum Images to track the progression of the process in the microscope.

Below are descriptions of the two scripts I need and I'm wondering if anyone may have scripted
something like this in the past, has an idea of how the script can be written or has alternative
means of achieving the same end result, within Digital Micrograph. I pose the question to the
listserver as I'm not much of a programmer/scripter and need all the help I can get.

Script 1:
I would like to record EELS spectra with the beam stationary on the specimen. I would like to do
this automatically with a script. The user, sets all acquisition parameters (in Digital Micrograph)
and also the number of spectra required and the time delay between the spectra (in the script). The
script starts, controls the acquisition, records all the spectra into a 'spectrum stack', saves the
output in a file to disk and returns control to the user. In the Results window the script outputs
the duration of the complete acquisition and if possible the duration of each spectrum acquisition
within the 'spectrum stack' or this information is saved as metadata/tags within the 'spectrum
stack' data file.

Script 2:
I would also like to do the same with Spectrum Imaging.
In this case the user sets up the Spectrum Imaging acquisition parameters (in Digital Micrograph)
and also the number of Spectrum Images required and the time delay between the Spectrum Images (in
the script). The script controls the acquisition and saves the EELS Spectrum Image, ADF Image and
the Survery Image with a logical numbering system and returns control to the user. In the Results
window the script outputs the duration of the complete acquisition and if possible the duration of
each spectrum image or this information is saved as metadata/tags within the 'spectrum image' data
files.
In the case where the time delay between the spectrum images is greater than the total acquisition
time of the spectrum image, I need Digiscan to continue scanning the probe over the defined ROI of
the spectrum image.
The script may require a warning/check to ensure the delay entered is an integer multiplier of the
acquisition time, so that the probe scans the full ROI before the next Spectrum Image is acquired.
If the probe scanned a proportion of the Spectrum Image ROI prior to the next Spectrum Image
acquisition that would be a big problem.

Any help that you can provide with the scripts would be greatly appreciated.
If you can write the scripts I can promise a few bottles of Australian wine, the adoration and
acclaim of your peers and my gratitude.

Best Regards
George

PS Does anyone have an email for Dr David Mitchell? The ANSTO email has failed. Please respond
directly to me. Many thanks.

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HI,

We have a 4pi EDS system that uses a PCI card which only runs in a Mac G4. The Mac is dying. I am looking for a 4pi SEII version 2.2 PCI card to run our EDS system. Unlike earlier models, that particular PCI card version will operate in a Windows PC.

We will be happy to buy this card. Please contact me offline if you have one of these cards that isn't being used.

Thanks,

Owen

Owen P. Mills
Materials Science & Engineering
Michigan Technological University
1400 Townsend Drive
Houghton, MI 49931
PH 906-369-1875
FAX 906-487-2934

==============================Original Headers==============================
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Organization: CIBNOR

Title-Subject: [Filtered] Theoretical and Practical Course

Message: Events
BIOLOGICAL RESEARCH CENTER NORTHWEST, SC,

THROUGH

Electron Microscopy Laboratory

Is pleased to invite you to:

"Basic Course SCANNING ELECTRON MICROSCOPY"

Location: Room 1, Building "S" and Electron Microscopy Laboratory

From 12 to 16 November 2012

La Paz, Baja California Sur, Mexico

General Coordination:

Arturo Cruz Ariel Tec Villacorta
CIBNOR, S.C.
52 + (612) 123-8484 Ext 3773

Contact:
Paulina Meza Navarro
Events Department
CIBNOR, S.C.
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E mail: microscopia2012-at-cibnor.mx

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Name: Ariel Cruz

Organization: CIBNOR

Title-Subject: [Filtered] Curso teorico practico

Message: Eventos
EL CENTRO DE INVESTIGACIONES BIOL�GICAS DEL NOROESTE, S.C.,

A TRAV�S DEL

Laboratorio de Microscopia Electr�nica

Se complace en invitarle al:

"CURSO B�SICO DE MICROSCOP�A ELECTR�NICA DE BARRIDO"

Lugar: Sala 1, edificio "S" y Laboratorio de Microscopia Electr�nica

Del 12 al 16 de noviembre del 2012
La Paz, Baja California Sur, M�xico

Coordinaci�n General:

Tec. Ariel Arturo Cruz Villacorta
CIBNOR, S.C.
52+ (612) 123-8484 Ext. 3773

Contacto:
Paulina Meza Navarro
Departamento de Eventos
CIBNOR, S.C.
52+ (612) 12 384-84 Ext. 3130
E mail: microscopia2012-at-cibnor.mx

--------------------------------------------------------------------------------

�ltima Actualizaci�n: Martes, 23 Agosto 2011, 12:00 - Silvia Yolanda Alzaga Mayagoitia

Instituto Polit�cnico Nacional 195, Playa Palo de Santa Rita Sur; La Paz, B.C.S. M�xico; C.P. 23096,
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Email: ravi.thakkar369-at-gmail.com
Name: Ravi Thakkar

Title-Subject: [Filtered] section folding in Cryo Ultramicrotomy

Message: Dear Listener,

How to prevent section folding in Cryo Ultramicrotomy..??

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A postdoctoral positions is available at Northwestern University to
work on Solid Oxide Fuel Cells. The successful candidate will work as
part of a team of PhD students, postdocs and faculty designing new
materials for extended life at lower temperatures. The work will
involve many different types of characterization ranging from SEM
through classic BF/DF to more advanced analytical or HREM � follow the
science not the electron. A strong general background in electron
microscopy as well as a good background in materials science and/or
SOFCs is essential, as are good writing and communication skills.
Ambition and drive to publish 4-6 papers a year is also important.

Interested applications should contact (by email) Prof L. D. Marks,
L-marks at northwestern.edu providing an electronic copy of their CV
with the names of three referees.

--
Laurence Marks
Department of Materials Science and Engineering
MSE Rm 2036 Cook Hall
2220 N Campus Drive
Northwestern University
Evanston, IL 60208, USA
Tel: (847) 491-3996 Fax: (847) 491-7820
email: L-marks at northwestern dot edu
www.numis.northwestern.edu

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Fellow Listers:

I'm looking for tips from those familiar with Adobe Photoshop's "Photomerge" tool. �
Has anyone used this for stitching multiple microscope field images, followed by subsequent quantitative image analysis?

I tried this feature for the first time yesterday and found that the image magnification calibration changed after merging.
Specifically a 100um scale bar placed on one of the original images was measured by Photoshop's ruler at ~200um on the resulting merged image.

Of course this is not a problem for a handful of images, we would just recalibrate. �
However we have a large volume to process in this way, and are wondering if anyone has gone through the steps to see how reproducible the outcome is? �
If one always begins with image sets captured on the same microscope, same objective, does the merged image always result in the same reproducibly changed pixel to micron ratio? �
Does this depend on the number of images being merged?
I would of course run the tests myself but this is a short-term project on a tight timeline.

I am also curious if anyone has looked into whether there are any spatial alterations following the merge. �
Visually the result is impressive, however there may be more than meets the eye?

Thanks in advance for any suggestions!

Karen

Karen Zaruba
University of Minnesota
Minneapolis, MN 55455

kszaruba-at-umn.edu

==============================Original Headers==============================
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How was the image calibrated and how was the measurement done? It would probably to help to know which software was used. I don't think that Photoshop enters into this, per se.

Some systems store the calibration as the dimensions of each pixel. That should not change with stitching. However, what should be the case and what is actually the case are often enough different. It is good to verify.

Other systems will calibrate based on the field width at a given magnification regardless of the number of pixels in the image. (We have one). If I took four images at 1000x with a 120-um horizontal field width and stitched them together, the results would be (nearly) twice as many pixels in each direction and would represent twice the view (240um). But 240 um is the field of view of a 500x image on our system. If I entered the mag of my new image as 1000x and measured the feature, I would be about 2x in error. A 100-um scale bar would be measured as 200 um.

It seems like that could be going on in your case. I would investigate that avenue.

Regards,
Warren

-----Original Message-----
X-from: kstzar-at-yahoo.com [mailto:kstzar-at-yahoo.com]
Sent: Wednesday, October 10, 2012 10:48 AM
To: wesaia-at-iastate.edu

Fellow Listers:

I'm looking for tips from those familiar with Adobe Photoshop's "Photomerge" tool. �
Has anyone used this for stitching multiple microscope field images, followed by subsequent quantitative image analysis?

I tried this feature for the first time yesterday and found that the image magnification calibration changed after merging.
Specifically a 100um scale bar placed on one of the original images was measured by Photoshop's ruler at ~200um on the resulting merged image.

Of course this is not a problem for a handful of images, we would just recalibrate. �
However we have a large volume to process in this way, and are wondering if anyone has gone through the steps to see how reproducible the outcome is? �
If one always begins with image sets captured on the same microscope, same objective, does the merged image always result in the same reproducibly changed pixel to micron ratio? �
Does this depend on the number of images being merged?
I would of course run the tests myself but this is a short-term project on a tight timeline.

I am also curious if anyone has looked into whether there are any spatial alterations following the merge. �
Visually the result is impressive, however there may be more than meets the eye?

Thanks in advance for any suggestions!

Karen

Karen Zaruba
University of Minnesota
Minneapolis, MN 55455

kszaruba-at-umn.edu

==============================Original Headers==============================
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12, 37 -- Subject: Image Analysis: Resolution changes after Photoshop Photomerge
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23, 38 -- From wesaia-at-iastate.edu Wed Oct 10 11:35:23 2012
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23, 38 -- From: "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu}
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23, 38 -- "Microscopy-at-microscopy.com"
23, 38 -- {Microscopy-at-microscopy.com}
23, 38 -- Subject: RE: [Microscopy] Image Analysis: Resolution changes after Photoshop
23, 38 -- Photomerge
23, 38 -- Thread-Topic: [Microscopy] Image Analysis: Resolution changes after
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Listers,

Thank you for the quick replies! �A number of you asked for more detail on the version of Photoshop being used (CS5 for Mac) and the steps for calibration (micrometer slide image, Photoshop Analysis } Set Measurement Scale } Custom).

Looking further into the details I realized somewhat sheepishly that I was unable to reproduce the error that we saw yesterday. �
Possibly we neglected to check the "Use Measurement Scale" box in Photoshop's Ruler menu bar on one image but not the other.
In any case the magnification error seems to have been one of operator origin.

Spatial alteration concerns remain: �I can't answer whether Photohop's algorithm involves simple translating of images or whether any stretching is going on. �I had hoped someone else might have explored this? �
If not,�these concerns will hopefully be dispelled by a few checks, such as measuring features "before" and "after" in portions including those where the images overlap.
�
Also thanks for the suggestion of Fiji J's�Track2EM option, which I may want to explore for another project!

Sincerely,

Karen Zaruba

----- Original Message -----
X-from: "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu}
To: "kstzar-at-yahoo.com" {kstzar-at-yahoo.com} ; "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
Cc:
Sent: Wednesday, October 10, 2012 11:35 AM

Fellow Listers:

I'm looking for tips from those familiar with Adobe Photoshop's "Photomerge" tool. �
Has anyone used this for stitching multiple microscope field images, followed by subsequent quantitative image analysis?

I tried this feature for the first time yesterday and found that the image magnification calibration changed after merging.
Specifically a 100um scale bar placed on one of the original images was measured by Photoshop's ruler at ~200um on the resulting merged image.

Of course this is not a problem for a handful of images, we would just recalibrate. �
However we have a large volume to process in this way, and are wondering if anyone has gone through the steps to see how reproducible the outcome is? �
If one always begins with image sets captured on the same microscope, same objective, does the merged image always result in the same reproducibly changed pixel to micron ratio? �
Does this depend on the number of images being merged?
I would of course run the tests myself but this is a short-term project on a tight timeline.

I am also curious if anyone has looked into whether there are any spatial alterations following the merge. �
Visually the result is impressive, however there may be more than meets the eye?

Thanks in advance for any suggestions!

Karen

Karen Zaruba
University of Minnesota
Minneapolis, MN 55455

kszaruba-at-umn.edu

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Dear Karen,

The Photomerge dialogue describes what changes it will do with the
pictures under each option: Auto, Perspective, cylindrical, spherical, and
collage will all scale images independently to get the best match (and
most of those options will do much worse distortions too).

The reposition option will not scale images. Do not select geometric
distortion correction, as if the objective is high quality there should be
next to none, and this process may introduce some.

Having used this feature extensively, I have learnt that the process is
not the same each time it is applied to the same set of images. If you see
a misalignment of images, or wrongly placed image, it is often possible to
re-run the process, and it will do the merge differently the next time.

Best regards,

Ben

--
Research Support Manager
MRC Anatomical Neuropharmacology Unit, Mansfield Road,
Oxford, OX1 3TH, United Kingdom. Telephone: 01865 271649
{http://www.mrc.ox.ac.uk/}

On 10/10/2012 23:06, "kstzar-at-yahoo.com" {kstzar-at-yahoo.com} wrote:

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The resolution issue in Photoshop could be due to either the method of
making the measurement or, possibly, the setting for the image cache. I
don't know whether or not the geometric methods for image stitching in
Photoshop contribute, since I only use Reposition and Interactive methods.

If you use the measure tool, the zoom of the image is tantamount to
getting an accurate measurement. If the image view is at less than
100%, then you will likely get the wrong measurement. The measure tool
appears to be tied to the screen resolution, and so the only accurate
method is to fill the screen with the scale bar and then draw with the
measure tool to get an accurate reading.

The same is true when drawing a line to create a scale bar. The reading
on the Info box is not accurate unless zoomed in.

A sure fire way to obtain an accurate measurement, no matter what the
zoom, is through the use of any means that would create a Region of
Interest or, in Photoshop, and outline that is referred to as "marching
ants." The magic wand tool or the Color Range tool with a Fuzziness set
at zero (0) can be used to create regions of interest. The reading from
the Info Box will then provide an accurate measurement.

That is a relief because the Analysis Tool in Photoshop requires regions
of interest.

As an added safety precaution, you might also want to set the memory
cache to 1 (versus the default setting). In Edit (PC) or Photoshop
(Mac) } Preferences, find the cache setting (in Performance on CS3) and
set this to 1 (one). If the cache is set higher, then Photoshop retains
low resolution copies of the image for display at lower zooms to save on
memory. I haven't really been convinced that a setting of 1 prevents
caching, but I'm more comfortable with this setting. I'll write to
Adobe to get more info.

I created a scale bar and put it through the paces to replicate the
scale bar differences when using "Reposition Only" as the photostitching
method. I could only get a 2 pixel difference in reading depending on
zoom when using the Measure tool. I'm on a PC.

Hope this helps,

Jerry

On 10/10/2012 11:41 AM, wesaia-at-iastate.edu wrote:
} ----------------------------------------------------------------------------
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}
} How was the image calibrated and how was the measurement done? It would probably to help to know which software was used. I don't think that Photoshop enters into this, per se.
}
} Some systems store the calibration as the dimensions of each pixel. That should not change with stitching. However, what should be the case and what is actually the case are often enough different. It is good to verify.
}
} Other systems will calibrate based on the field width at a given magnification regardless of the number of pixels in the image. (We have one). If I took four images at 1000x with a 120-um horizontal field width and stitched them together, the results would be (nearly) twice as many pixels in each direction and would represent twice the view (240um). But 240 um is the field of view of a 500x image on our system. If I entered the mag of my new image as 1000x and measured the feature, I would be about 2x in error. A 100-um scale bar would be measured as 200 um.
}
} It seems like that could be going on in your case. I would investigate that avenue.
}
} Regards,
} Warren
}
} -----Original Message-----
} X-from: kstzar-at-yahoo.com [mailto:kstzar-at-yahoo.com]
} Sent: Wednesday, October 10, 2012 10:48 AM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] Image Analysis: Resolution changes after Photoshop Photomerge
}
}
}
}
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}
} Fellow Listers:
}
} I'm looking for tips from those familiar with Adobe Photoshop's "Photomerge" tool.
} Has anyone used this for stitching multiple microscope field images, followed by subsequent quantitative image analysis?
}
} I tried this feature for the first time yesterday and found that the image magnification calibration changed after merging.
} Specifically a 100um scale bar placed on one of the original images was measured by Photoshop's ruler at ~200um on the resulting merged image.
}
} Of course this is not a problem for a handful of images, we would just recalibrate.
} However we have a large volume to process in this way, and are wondering if anyone has gone through the steps to see how reproducible the outcome is?
} If one always begins with image sets captured on the same microscope, same objective, does the merged image always result in the same reproducibly changed pixel to micron ratio?
} Does this depend on the number of images being merged?
} I would of course run the tests myself but this is a short-term project on a tight timeline.
}
}
} I am also curious if anyone has looked into whether there are any spatial alterations following the merge.
} Visually the result is impressive, however there may be more than meets the eye?
}
} Thanks in advance for any suggestions!
}
} Karen
}
} Karen Zaruba
} University of Minnesota
} Minneapolis, MN 55455
}
} kszaruba-at-umn.edu
}
}
}
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} 23, 38 -- From: "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu}
} 23, 38 -- To: "kstzar-at-yahoo.com" {kstzar-at-yahoo.com} ,
} 23, 38 -- "Microscopy-at-microscopy.com"
} 23, 38 -- {Microscopy-at-microscopy.com}
} 23, 38 -- Subject: RE: [Microscopy] Image Analysis: Resolution changes after Photoshop
} 23, 38 -- Photomerge
} 23, 38 -- Thread-Topic: [Microscopy] Image Analysis: Resolution changes after
} 23, 38 -- Photoshop Photomerge
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--
Jerry (Gerald) Sedgewick
Scientific Imaging & Image Analysis Consultant
Technical Writer / Regulatory Consultant
965 Cromwell Avenue
Saint Paul, MN 55114
651-788-2261
http://www.imagingandanalysis.com
http://www.quickphotoshop.com
Author: �Scientific Imaging with Photoshop: Methods, Measurement and Output�

==============================Original Headers==============================
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-----Original Message-----
X-from: Steven Samuelsson
Sent: Thursday, October 11, 2012 9:52 AM
To: Margo Gill-Linscott

The Histochemical Society Immunohistochemistry and Microscopy: A
Hands-on Course, First Announcement
March 9 - 13, 2013, Marine Biological Laboratory, Woods Hole,
Massachusetts USA

Colleagues,

The Histochemical Society, in partnership with the Marine Biological
Laboratory, is offering our sixth annual Immunohistochemistry and
Microscopy Hands-on Course, on the campus of the Marine Biological
Laboratory at Woods Hole, Massachusetts. The course begins on the
evening of March 9, (students should plan to arrive at MBL on March 9)
and ends at the end of the day on March13. Departure from MBL will be on
March 14.

The course is four days of in-depth theory of and extensive hands-on
experience with immunohistochemistry (IHC) techniques as well as theory
and hands-on experience with a broad range of microscopic imaging
techniques. The course taught by experienced academic researchers and
knowledgeable industry representatives, allows for close interaction
between faculty and students and small group discussions.

Course Topics:
Antigen retrieval
Automated immunohistochemistry
Chromogenic detection
Controls
Double labeling and co-localization
Fixation
Fluorescence detection
Microscopy and imaging
Troubleshooting.

Registration includes accommodations and meal package.

Please visit the Immunohistochemistry and Microscopy website for more
information: http://immunohistochem.com/

Meg McGough
The Histochemical Society

==============================Original Headers==============================
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9, 20 -- - 1st Announcement
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Please share

Advanced Microscopy LIVE Workshop
January 14-18, 2013
University of California, Santa Barbara

Offered by the Neuroscience Research Institute (NRI) and Department of
Molecular, Cellular, and Developmental Biology (MCDB).
See: https://www.lifesci.ucsb.edu/mcdb/events/imaging-live/

This workshop is five days of seminars and hands on instruction. Through
the course, participants will gain theoretical and practical experience
with light microscopy on live specimens. Lectures will cover basic to
advanced microscopy principals with an emphasis on live imaging
applications.

--
Mary Raven
Microscopy Facility Director
Neuroscience Research Institute &
Molecular, Cellular & Developmental Biology
Biology 2 Building, room 5173
The University of California
Santa Barbara, CA 93106-5060

Vacation Dates
Thanksgiving 2012 Nov 21- Nov 25
Winter Break 2012 Dec 21-Jan 6

http://www.lifesci.ucsb.edu/~m_raven/
Phone: (805) 893 8702
Fax: (805) 893 2005

==============================Original Headers==============================
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9, 20 -- Microscopy LIVE Workshop
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Hi Folks,

We need to replace the HT-cable on our JEOL 8600 electron microprobe.
If you have one,
or know of anyone who has one available, please contact me offline.
We'll gladly pay for it.

Chris

--
Chris Fleisher
Electron Microprobe Lab Coordinator
Department of Geology
University of Georgia
Athens, Ga. 30602
Ph: (706) 542-2419 E-mail: fleisher-at-gly.uga.edu

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Hi Chris

If all else fails look up an organisation that deals with x-ray equipment
for medial or industrial applications as in my past, in several countries,
they have been able to produce an excellent cable just by me supplying the
cable ends. I should also say they have been more economical than the
original equipment manufacturer!

Good luck

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
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Sent: 12 October 2012 14:08
To: protrain-at-emcourses.com

Hi Folks,

We need to replace the HT-cable on our JEOL 8600 electron microprobe.
If you have one,
or know of anyone who has one available, please contact me offline.
We'll gladly pay for it.

Chris

--
Chris Fleisher
Electron Microprobe Lab Coordinator
Department of Geology
University of Georgia
Athens, Ga. 30602
Ph: (706) 542-2419 E-mail: fleisher-at-gly.uga.edu

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Dear List,
I have some ideas for improving thermal and electrical conductivities
at LHe temperatures that I'd like to study. In order to do this I
need both financial and personnel support. An undergraduate in the
lab at Caltech and I have done some preliminary experiments, which
showed promise. Specimen preparation is easy, and I can arrange to
use a facility on which to do the experiments--there will be a charge
for it. I would insist that the undergraduate who helped with the
preliminary experiments be a co-author on at least the first
publication. Please contact me off-line if you would be able to
support this project, and I'll let you know the details.
Yours,
Bill

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Recently, in the middle of the night I found myself, not counting sheep,
but trying to solve in my head the following: For a three-dimensional
lattice with primitive base vectors a, b, c, find a vector perpendicular
to the bc plane and with a length equal to the spacing of those planes.
(Here and throughout, please imagine that a, b and c are in bold to
represent vectors.) I have no idea why I wanted to know this; perhaps I
needed it in the dream from which I awoke. (Note that the vector sought
is not the reciprocal lattice vector a* since that has a length that is
not the plane spacing but the reciprocal of the plane spacing.) The
result I got was (derivation below):

(a.b�c)b�c/(b�c).(b�c)

Now this expression is more complicated than I anticipated. So my
question is: can anyone tell me of a simpler expression?

P.S. It did not help me get back to sleep.

Derivation: We need a vector perpendicular to the bc plane. b�c is
such a vector. Thus a unit vector in that direction is b�c/|b�c|. The
length of the vector needs to be the plane spacing which is the
projection of the vector a onto this unit vector, namely a.b�c/|b�c|.
The final result is obtained by multiplying the unit vector in the
appropriate direction by the (scalar) length of the vector to give the
result above.

Alwyn Eades
--
Alwyn Eades
Department of Materials Science and Engineering
Lehigh University
610 758 4231

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Hello everyone,

We are keen to locate facility that can provide FEI Osiris TEM services.
Our interest is 2D dopant profiling in silicon.

Would anyone know an outside lab, university, or industrial concern
that might offer these services?

with best regards,

bryan tracy

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Dear colleagues,
�
I am afraid I am not able to solve a seemingly basic problem.
We have an inverted Zeiss light microscope with a ccd camera (PixelLink) just above the occulars.
There is a lense (a ring) between the camera and the microscope.
We have 3 fix halos on the background and I can't find the origin of the dust. The halo appears as if the dust were near focus.
Here are the tests I did in order to diagnose the problem:
- When I turn the camera with the ring attached (both turn together in relation to the microscope), the halos don't move even a bit.
--} the dust must be part of the camera/ring, not the microscope.
�
- If I detach the camera from the microscope (with the ring attached), the background (when I direct the camera to the light coming from the window) is perfectly clean.
--} the dust are NOT part of the camera/ring but are part of the microscope ?!!
�
It makes no sense! Of perhaps the ambiant light is not enough to give a clear picture?? (I have a bright uniform background though)
�
One additional note: if I make the ring loose and I slowly shake�the camera/ring a bit, the halos shake also a bit on the picture.
I already cleaned the lense/ring and the fine glass in front of the camera CCD with lense paper and ethanol.
Any comment welcome.
�
Best regards,
Stephane

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2, 37 -- From: Stephane Nizet {nizets2-at-yahoo.com}
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2, 37 -- Subject: Cleaning an LM microscope with CCD camera
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Dear Stephane,

} - If I detach the camera from the microscope (with the ring attached),
} the background (when I direct the camera } to the light coming from the
} window) is perfectly clean.
} --} the dust are NOT part of the camera/ring but are part of the
} microscope ?!!

The light coming through the microscope is coming at the camera/coupling
optics from a small range of angles, making the dust cast sharp shadows on
the sensor. Off the microscope the light comes into the camera/coupling
optics from a wide range of angles, and the shadows are softened to the
point of being invisible. The same happens with dust on DSLR sensors-
often dust is not visible until you stop down a lens to f/11 or so. You
could try performing the same test in a dark room with a small
high-powered LED torch located a few feet away.

When you say you cleaned the glass in front of the CCD, do you mean an IR
(hot mirror) filter in front of the sensor? If so, there could be dust on
the internal side of the hot mirror (if there is a significant gap between
that and the sensor). Does the hot mirror have a retaining ring allowing
its removal?

Regards,

Ben

On 15/10/2012 12:43, "nizets2-at-yahoo.com" {nizets2-at-yahoo.com} wrote:

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Greetings,

Today is the last day to register at only $25 for the full day Workshop on Biological structures and techniques on November 14th.

Info: http://mrl.illinois.edu/news/conferences-workshops/biological-structures-electron-optical-imaging-workshop

Registration is still open after today, but at $35.

This is a great workshop for those in material sciences to get a chance to look at biological structures, and for those learning about electron microscopy in the biological studies.

Lou Ann Miller
CMM Staff/ Frederick Seitz Material Research Laboratory.

==============================Original Headers==============================
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Email: j.janssen-at-nki.nl
Name: Hans Janssen

Organization: Netherlands Cancer Institute

Title-Subject: [Filtered] Balzers CTA010

Message: Dear listers,
We had a meltdown of the carbon evaporation lid from our Balzers CTA010 (glow discharge/carbon
evaporator). Since the machine is very old there are no parts readily available. Is there anybody
out there that could provide us such a lid, maybe from a machine that is kept in the dungeons of
your institute?
Thanks in advance, Hans.

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Dear listers,
�
First of all I want to thank those who responded to my question about cleaning our PixeLink�camera.
Again I learned something, even if I was unable to solve the problem. It seems that the dust is firmly attached to the chip and I don't want to risk a harsh handling for 3 halos. We'll probably just substract a background image as a temporary solution but unfortunately the simple capture software doesn't do it automatically and in the long term I don't think that we can do the substraction with photoshop for each picture.
The camera is old (in terms of CCD camera) and its resolution is limited. We will consider the purchase of a new basic camera for the work.
The camera must be color, with a resolution of 3-5 Mpixel. It is for basic documentation at�magnifications from 20-60x. We already have a c-mount 0,63x adapter from Zeiss so ideally the camera should fit in.
Please share your advise with me with an approximate price.
�
Second question, this one more personal: I am the proud father of 2 wonderful children (this is completely subjective) and as a microscopist I would naturally like to share my passion for microscopy with them. I would like to offer them a simple light microscope for christmas. Not a toy but still a�basic one. Perhaps a stereomiscroscope would be a good idea because the objects wouldn't have to be flat and light transparent. I know there are some "new" USB microscopes�of good quality on the market but I would need advices or personal experiences and opinions to guide me.
Many thanks in advance (I thank you all in advance so I don't need to get 20 out-of-office replies just to thank you afterward).
�
Stephane

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2, 37 -- Subject: basic Microscope camera - advice
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} Name: Hans Janssen
}
} Organization: Netherlands Cancer Institute
}
} Title-Subject: [Filtered] Balzers CTA010
}
} Message: Dear listers,
} We had a meltdown of the carbon evaporation lid from our Balzers
} CTA010 (glow discharge/carbon
} evaporator). Since the machine is very old there are no parts
} readily available. Is there anybody
} out there that could provide us such a lid, maybe from a machine
} that is kept in the dungeons of
} your institute?
} Thanks in advance, Hans.

Dear Hans,
In case no one has a replacement lid, would it be possible to make
one with a 3D printer?
Yours,
Bill

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On Oct 16, 2012, at 12:47 AM, nizets2-at-yahoo.com wrote:

} Second question, this one more personal: I am the proud father of 2
} wonderful children (this is completely subjective) and as a
} microscopist I would naturally like to share my passion for
} microscopy with them. I would like to offer them a simple light
} microscope for christmas. Not a toy but still a basic one. Perhaps a
} stereomiscroscope would be a good idea because the objects wouldn't
} have to be flat and light transparent. I know there are some "new"
} USB microscopes of good quality on the market but I would need
} advices or personal experiences and opinions to guide me.
} Many thanks in advance (I thank you all in advance so I don't need
} to get 20 out-of-office replies just to thank you afterward).

Dear Stephane,
Project Micro on the MSA web site has a lot of info about this.
Yours,
Bill

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Email: j.knowles-at-ucl.ac.uk
Name: Jonathan Knowles

Organization: University College London

Title-Subject: [Filtered] Electronics side of an EDX system

Message: Hi

We have inherited an EDX system which seems to work OK (sorry I don't have details to hand but
wanted to ask this before I forgot!). The system works OK, but the electronics side is really
ancient (It has a microVAX type drive with about 20Mb storage to run the OS and software) and am
concerned that it might die some time. I wondered if anyone had ever built some more up to date
hardware/interface? The connections for the EDX consist of a BNC for the HT and then a 9 pin
connector presumably for data.

Thanks

Jonathan

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Dear Listers,
is there anybody out there who can point me to a source which kind of heater plates are used in the JEOL 6400 diffusion pumps?
Anybody out there who has heater plates and can part wih them for a reasonable price?
I am trying to revive this SEM for a public charity in Italy...

Best regards,
Stefan

--

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

==============================Original Headers==============================
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Greetings Fellow Microscopists,

The Midwest Microscopy and Microanalysis Society (M3S) will mark our 55th
anniversary with a meeting on Thursday, November 15, 2012 at Baxter
Healthcare, Deerfield, IL, followed by dinner at Ristorante Abruzzo,
Deerfield, IL.

"Microscopy For Life: M3S Celebrates 55 Years" is being organized by Robb
Mierzwa (Mierzwa-at-jeol.com) and Elaine Schumacher
(eschumacher-at-mccrone.com). The program includes presentations by MAS Tour
Speaker John Henry Scott, NIST, MSA Tour Speaker Janet H. Woodward, Buckman
Laboratories, Inc., Nestor Zaluzec, Argonne National Laboratory, and Scott
Stoeffler, McCrone Associates. In addition there will be presentations from
the six students who submitted their M&M 2012 abstracts for consideration
for M3S travel awards.

Program details and registration information can be found on the Meetings
page of our website:

http://www.midwestmicroscopy.org/meetings.htm

Please join us for our final meeting of 2012. We look forward to seeing
you there!

Elaine Schumacher
M3S Program Coordinator

Alan W Nicholls, PhD
Associate Director, RRC
Director of Research Service Facility (Electron Microscopy)
Research Resources Center - East (M/C 337)
Room 110 Science and Engineering South Building
The University of Illinois at Chicago
845 West Taylor St
Chicago, IL 60607-7058

Tel: 312 996 1227
Fax: 312 996 8091
Office: Room 110

Web site www.rrc.uic.edu

==============================Original Headers==============================
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As a part of our Chapter 11 restructuring, my employer has exited a
business and has a surplus Cambridge S360 SEM. Our first preference is to
see if here is someone who would like the instrument either for direct use
or for parts. The instrument was purchased around 1998 and was used in my
lab until 2003 for particle sizing. The microscope can use LaB6 cathodes
and has a digital imaging system (Olympus-SIS ADDA-II external scan
Interface and AnalySIS Pro Image processing software that runs under
Windows XP.) The microscope has limited remote capabilities - it was a
first generation remote interface and we could aid the operator but not
implement robust, unattended operation. The microscope also has a STEM
detector (our preference for particle sizing.) The scope has been used
intermittently since 2009 and is currently off and under vacuum. It worked
the last time it was used. It was quite robust while in my lab. It's last
PM was in 2009. The last service was through Leica. I have not really
followed the microscope since we passed it on, so I cannot speak to the
availability of parts.

If someone wants the instrument, they would be responsible for removing it.
My management would like it removed by the end of November (we need to
vacate the building.) Otherwise, it will be recycled. If someone has
serious interest, please respond to my work email: john.r.minter-at-kodak.com. (I
normally don't have time for the listserver at work and so am subscribed
with my home account.) I can help get you connected to the right people to
streamline the paperwork. I'm sure most of my fellow microscopists
understand -- we get attached to these instruments and prefer to see them
used - either directly or indirectly to keep another one going - rather
than be recycled.

Best Regards,
John

John Minter
Microscopy and Surface Science Laboratory
Kodak Analytical Sciences
Rochester, NY
john.r.minter-at-kodak.com

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Email: vakimler-at-med.wayne.edu
Name: Vickie A. Kimler

Organization: Wayne State University School of Medicine

Title-Subject: [Filtered] Fluoronanogold immunolabeling of isolated fat droplets

Message: In my hands, it appears that
LR-White is not amenable to embedment of isolated
lipid droplets for Fluoronanogold
immunolabeling of protein antigens on lipid droplets.

Does anyone know of a good embedding medium for isolated organelles (sucrose density gradient
purified) that would allow for good post embedding immunolabeling of these droplets on sections of
these droplets?

Additional comment: We want to also run electron tomography on these images - will the embedding
resin you suggest be amenable for tomography (IVEM)?
Thanks!

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Hi, Vickie,

It's been my experience in working with phospholipids in the lung surfactant system, immunogold labeling proteins associated with these lipids, that I needed to resort to Lowicryl HM20 embedding resin, and use freeze-substitution techniques to preserve the lipids and get good labeling. LR White resin dissolved phospholipids. It's been decades since I did my work, but my paper is in J Histochem Cytochem 40(10):1491-1500, 1992 http://jhc.sagepub.com/content/40/10/1491.full.pdf+html . The non-polar nature of the HM20 embedding resin favors the preservation of the lipids. Processing samples at low temperature after stabilizing the lipids with uranyl acetate and dehydrating my samples with methanol, which was less reactive with the phospholipids than acetone, helped to preserve the phospholipids in the lung tissue and extracellular airways of the rat lung I was studying for my immunogold labeling experiments. The paper is over 20MB in size, which is why I did not attach it to this message.

Ed Haller

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
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Email: vakimler-at-med.wayne.edu
Name: Vickie A. Kimler

Organization: Wayne State University School of Medicine

Title-Subject: [Filtered] Fluoronanogold immunolabeling of isolated fat droplets

Message: In my hands, it appears that
LR-White is not amenable to embedment of isolated
lipid droplets for Fluoronanogold
immunolabeling of protein antigens on lipid droplets.

Does anyone know of a good embedding medium for isolated organelles (sucrose density gradient
purified) that would allow for good post embedding immunolabeling of these droplets on sections of
these droplets?

Additional comment: We want to also run electron tomography on these images - will the embedding
resin you suggest be amenable for tomography (IVEM)?
Thanks!

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23, 44 -- "Microscopy-at-microscopy.com" {Microscopy-at-microscopy.com}
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Email: mulqueen-at-gatan.com
Name: Joe Mulqueen

Organization: Gatan Inc

Title-Subject: [Filtered] Digital Micrograph menu bar bug

Message: This is inresponse to the problem of a DM window getting stuck undr the menu bar,submitted
by Se Wee Chee on 9/27.

. When the image gets stuck under the menu bar click and drag on the side of the image as if you
are making it smaller. Then when you let go of the mouse the window will be resized to a smaller
window and the top of the window will be visable. I hope this helps.

Sincerely
Joe Mulqueen
Gatan Inc

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Hi,
Just adding to the email of Joe: If you're unable to access a window by mouse interaction, you may access it by scripting.
(Open ScriptEditor "File/New Script..." enter the script and execute it either by the execute button, or by pressing CTRL & ENTER while the cursor is blinking in the script window.)

/*Script to shift the front most image 50 pixels downwards:*/

DocumentWindow win=GetFrontImageDocument().ImageDocumentGetWindow()
Number t,l,b,r
win.WindowGetFrameBounds(t,l,b,r)
win.WindowSetFrameBounds(t+50,l,b+50,r)

/* Script to change the minimize/maximize the document window: */

DocumentWindow win=GetFrontImageDocument().ImageDocumentGetWindow()
Number kNORMAL = 0
Number kMAXIMIZE = 1
Number kMINIMIZE = 2
Number kRESTORE = 3
win.WindowSetViewState(kNORMAL)

regards,
Bernhard

--------------------------------------------------------
Dr. Bernhard Schaffer,
Applications Software Developer
Gatan R&D
�
Office:+49 (0)89358084 70
Fax:�� +49 (0)89358084 77
bschaffer-at-gatan.com
http://www.gatan.com
--------------------------------------------------------

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Sent: 19 October 2012 02:44
To: Bernhard Schaffer

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Email: mulqueen-at-gatan.com
Name: Joe Mulqueen

Organization: Gatan Inc

Title-Subject: [Filtered] Digital Micrograph menu bar bug

Message: This is inresponse to the problem of a DM window getting stuck undr the menu bar,submitted by Se Wee Chee on 9/27.

. When the image gets stuck under the menu bar click and drag on the side of the image as if you are making it smaller. Then when you let go of the mouse the window will be resized to a smaller window and the top of the window will be visable. I hope this helps.

Sincerely
Joe Mulqueen
Gatan Inc

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Hi,
Just adding to the email of Joe: If you're unable to access a window by mouse interaction, you may access it by scripting.
(Open ScriptEditor "File/New Script..." enter the script and execute it either by the execute button, or by pressing CTRL & ENTER while the cursor is blinking in the script window.)

/*Script to shift the front most image 50 pixels downwards:*/

DocumentWindow win=GetFrontImageDocument().ImageDocumentGetWindow()
Number t,l,b,r
win.WindowGetFrameBounds(t,l,b,r)
win.WindowSetFrameBounds(t+50,l,b+50,r)

/* Script to change the minimize/maximize the document window: */

DocumentWindow win=GetFrontImageDocument().ImageDocumentGetWindow()
Number kNORMAL = 0
Number kMAXIMIZE = 1
Number kMINIMIZE = 2
Number kRESTORE = 3
win.WindowSetViewState(kNORMAL)

regards,
Bernhard

--------------------------------------------------------
Dr. Bernhard Schaffer,
Applications Software Developer
Gatan R&D

Office:+49 (0)89358084 70
Fax: +49 (0)89358084 77
bschaffer-at-gatan.com
http://www.gatan.com
--------------------------------------------------------

-----Original Message-----
X-from: microscopylistserver-noreply-at-microscopy.com [mailto:microscopylistserver-noreply-at-microscopy.com]
Sent: 19 October 2012 02:44
To: Bernhard Schaffer

X-from: mulqueen-at-gatan.com ()

This Question/Comment was submitted to the Microscopy Listserver using the WWW based Form at http://microscopy.com/MicroscopyListserver/MLFormMail.html
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Email: mulqueen-at-gatan.com
Name: Joe Mulqueen

Organization: Gatan Inc

Title-Subject: [Filtered] Digital Micrograph menu bar bug

Message: This is inresponse to the problem of a DM window getting stuck undr the menu bar,submitted by Se Wee Chee on 9/27.

. When the image gets stuck under the menu bar click and drag on the side of the image as if you are making it smaller. Then when you let go of the mouse the window will be resized to a smaller window and the top of the window will be visable. I hope this helps.

Sincerely
Joe Mulqueen
Gatan Inc

Login Host: 209.195.180.226
Listserver Email Form V - 20120416
---------------------------------------------------------------------------
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Dear listeners,

I tried to change by myself my FEG tip on a cold emission gun of a JEOL
6400F. Changing the tip itself cost only 1931.95 euros in France while
normal exchange procedure cost 16617.18 euros. Of course second option
is easier because tip is already mounted and centered on an assembly
which include insulator and extracting anode (whole assembly is
delivered and old one is send back to factory). My first test is rather
interesting because I succeed to instal the tip inside gun and to find
convenient procedure to do good conditionning of gun. Finally I
obtained good conditions of emission and image on the screen.
Unfortunately, as was afraid of it, only a few part of emission reach
bottom of the column. This is because mechanical centering of the tip
inside the assembly is critical. Tip should be very straight and well
and well centered with extracting anode hole.
I wanna try again to adjust tip mechanically and that the reason why I
ask for help of listeners for any idea to adjust tip mechanically and
particulary what kind of optical material could be convenient for such
work, how distance between tip and extracting anode is effective, etc...
Thanks a lot for your attention.
--
Nicolas STEPHANT

Universit� de Nantes
Institut Jean Rouxel
Service de microscopie �lectronique � balayage et microanalyse
2 rue de la Houssini�re
BP 92208
44322 Nantes c�dex 3

T�l : 02 40 37 64 26
M�l : nicolas.stephant-at-univ-nantes.fr
Web : http://www.univ-nantes.fr/smebm

"Le monde n'existe que pour autant que nous sommes capables d'en
produire une image"

C.G Jung

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Hello,

Does anyone have a manual for the Fullam EMS 76M sputter coater?

Or, else can answer some questions about it?

1) What's the difference in the electronics between etch and coat? Reversal in polarity + different voltage? What are the voltages?
2) What are the units for the vacuum gauge? What's the optimal pressure (Au in Ar).
3) When I change the current knob, that is adjusting the voltage slightly, right? I read the result on the current gauge which is the current between the anode an cathode?

Bonus: I notice nobody seems to use sputter coaters for carbon. Why?

Thank you in advance!

Zack Gainsforth
Space Sciences Laboratory, UC Berkeley
7 Gauss Way
Berkeley, CA 94720
510-642-9733
zackg-at-ssl.berkeley.edu

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12, 22 -- Subject: Fullam EMS 76M sputter coater manual/questions?
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Email: cmazareanu-at-yahoo.com
Name: Constantin

Title-Subject: [Filtered] old SEM

Message: Hi, I have an old Hitachi SEM and I want to bring it to digital age. I did buy a ADDA II
box. In order to connect this box to a computer is necessary a special pc board - Grabbit frame
grabber with dual fiber optic ports. This board is very old and cannot find it anywhere. Do you know
a solution/substitute for this frame grabber?
Thank you very much for your support!

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Hi Constatin, Point Electronic (http://www.pointelectronic.de/ ) do digital
upgrades of SEMs - you could see if they have something.... Good luck.

Ian
Ian Holton PhD
Acutance Scientific Ltd
1 Steel Bridge Cottages
Sham Farm Road
Eridge Green
Tunbridge Wells
TN3 9JD
Tel -+44 845 4796989
Mob +44 7774 394519
Skype acutance
Web www.acutance.co.uk
Registered at the above address, company number 7469246

Ian Holton
Acutance Scientific

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Email: cmazareanu-at-yahoo.com
Name: Constantin

Title-Subject: [Filtered] old SEM

Message: Hi, I have an old Hitachi SEM and I want to bring it to digital
age. I did buy a ADDA II box. In order to connect this box to a computer is
necessary a special pc board - Grabbit frame grabber with dual fiber optic
ports. This board is very old and cannot find it anywhere. Do you know a
solution/substitute for this frame grabber?
Thank you very much for your support!

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Organization: UT Southwestern Medical Center at Dallas

Title-Subject: [Filtered] EM Tech position available

Message: The Electron Microscopy Core Facility (EMCF) at UT Southwestern has a position open
immediately for an Electron Microscope Technician. Level of appointment and salary will depend on
qualifications and experience. The job posting can be found here:

sws001.swmed.edu/hr/jobDetail?postingNbr=13000448&jobCategory=FRESH

Apply online here:

https://sws001.swmed.edu/trm/TrmResume

Company profile:

UT Southwestern Medical Center, located in Dallas, TX, ranks among the top academic medical centers
in the world. Its faculty members, who are responsible for a broad array of groundbreaking
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Sciences. The EMCF is a fee for service, institutionally subsidized, core facility that provides
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facility website can be found here:

http://www4.utsouthwestern.edu/mcif/index.htm

Kate Luby-Phelps, Interim Director
Electron Microscopy Core Facility
UT Southwestern Medical Center
Dallas, TX

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Email: hooghan-at-verizon.net
Name: Bobby Hooghan

Organization: Weatherfordlabs

Title-Subject: [Filtered] Tilt Correction of Images obtained in FEI Helios 650 Dual Beam

Message: Hi all,
We are appealing to the collective Imaging knowledge/resources that we have available on this list
server.
We use an FEI Helios 650 Small Dual Beam (SDB) in order to acquire Slice and View data sets on Shale
samples. 3D volumes are then generated using Avizo Fire.
Our clients would like to get an equidemnsional pixel (ideally it would be an equidimensional
voxel). Data sets currently obtained, have different X Y dimensions for pixels, because of the
fact that we image the surface -at- a 45/52 degree angle.
The questions we have are two fold:
1)Is it possible to adjust for the tilt after obtaining the datasets using external Imaging S/W? If
so, how can we accomplish that?
2) There is an option for tilt correction on the SDB s/w where one can set it up for an automatic
tilt correction. Is there a method using a known standard (like a stack of semiconductor thin
films)that can be cross-sectioned in the SDB and measured with and without tilt correction to know
that the tilt correction is working as it should? We are quite aware that the FEI S/W was verified
when it was put on but having an additional piece of data to support the tilt correction claim would
not hurt,

Thanks in advance,
Best regards,
Bobby Hooghan

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A while back I posted a question about writing an expression for the
vector distance between planes in a crystal. I found the expression
(a.b�c)b�c/(b�c).(b�c)
and asked if anyone tell me of a simpler expression? I thought I would
do an update because a couple have people have sent me messages to say
that there is no simpler result, but Phil Ahrenkiel of South Dakota
School of Mines and Technology sent me the beautiful and simple result
a*/(a*.a*)
where a* is the reciprocal lattice vector for the crystal.

--
Alwyn Eades
Department of Materials Science and Engineering
Lehigh University
610 758 4231

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Hello everyone.

I am trying to help a student with EM immunolabeling of GFP in retinal
rod cells.

Our pre-embedding DAB approach met with only limited success.
Vibrotomed sections were too fragile to carry through all the labeling
steps, so we worked with a whole dissected retina. With saponin
permeabilization we only got penetration of the outer parts of the
rods. Freeze thaw did not seem to help the penetration, though we may
not have done enough cycles. TritonX100 worked better, but really
disrupted the cell structure.

We are now thinking of trying a post-embedding approach using either
LRGold and progressive lowering of temperature, or cryofixation with
HM20 and AFS freeze sub, both of which have been done here, but not
for GFP. Can anyone suggest a good protocol? I understand that we
should leave some water in the prep, but am not sure how much to add,
and whether to add it to both solvent and resin. Also, what solvent
is best for dehydration for GFP?

Any suggestions of procedures or papers would be most welcome.

Dr. Marie E. Cantino
Associate Professor of Physiology and Neurobiology
Director, Electron Microscopy Laboratory
University of Connecticut, Unit 3242
Phone: 860-486-3588
Fax: 860-486-6369

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Email: fionacxr-at-tll.org.sg
Name: Fiona Chia

Organization: TLL

Title-Subject: [Filtered] TEM AFS Machine and Liquid Nitrogen

Message: Hi all,

My department just got a Leica EM AFS machine. I would like to know if
there is any concerns or safety issue with leaving liquid nitrogen in
the AFS when the AFS is not in use. Does the AFS have to be emptied of
liquid nitrogen after each use if no one is going to use it for a few
days? Would there be any concerns about pressure build up in the dewar?

Would appreciate any advice on this.

regards
Fiona Chia
Microscopy Technologist, Temasek Lifesciences Laboratory
Email: fionacxr-at-tll.org.sg

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Email: exploratorium-at-tiscali.it
Name: Giovanni

Organization: Museo delle Scienze Montalbo' - Casalciprano - Italy

Title-Subject: [Filtered] JEOL 6400 SEM

Message: Hi all,

We have been donated a JEOL 6400 SEM for our teaching/divulgative activities toward youngsters (see
our website: http://web.tiscali.it/exploratorium). We seek for anyone who has experience with the
use of this SEM for hints about its deinstallation, transportation, reinstallation and use. Thanks a
lot.
Giovanni De Caro, MD
Italy

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Email: mark.grimson-at-ttu.edu
Name: Mark Grimson

Organization: Texas Tech University

Title-Subject: [Filtered] TEM imaging of SWNT and MWNT

Message: Hello. Could I get some help on microscope set-up and sample prep for CNTs? A researcher
wants to look at microwave effects on single and multiwalled CNTs (shape defects, end defects). I
was going to use thin holey carbon support films at 75KV, but have seen nice images of CNTs at
200kV. Any suggestions?

Also, do CNTs generate enough contrast on their own, or could negative staining be used to help
improve the fine surface detail required to observe or amplify the subtle defects I am supposed to
see. Thanks in advance for any suggestions, before I begin imaging.

Thanks, Mark Grimson
Texas Tech University

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Hello Mark,

I have, in the past, done extentive TEM work on both SWNT and MWNT of Carbon. I was able to very successfully use a 300KV FEG TEM beam (at 300kv and also at 200kv) to get a good contrast on CNTs (without any negative staining). A lot of HRTEM was performed with great results. Ofcourse the SWNTs are more challenging as they tend to damage (deform and eventually decompose) under the beam faster than the MWNTs. I however was able to get HRTEM on SWNTs by allowing exposure under the electron bean for the least amount of time and also by using low dose method/s.

I used grids with holey carbon or holey carbon/formvar support films. I would put the CNTs in a vial with appropriate suspension, very lightly sonicate and use a micro-pipette to put a drop on the grid. Let it naturally dry for a little bit and overnight it in a desiccator (with a clean vacuum pump attached) before take it to the TEM. In cases where suspension does not carry any oil in it or it is simply alcohol, a natural dry for an hour or two is good enough before TEM.

My 2 cents...

Zia ur Rahman
Johnson Space Center, NASA,
Houston, TX

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Organization: Texas Tech University

Title-Subject: [Filtered] TEM imaging of SWNT and MWNT

Message: Hello. Could I get some help on microscope set-up and sample prep for CNTs? A researcher wants to look at microwave effects on single and multiwalled CNTs (shape defects, end defects). I was going to use thin holey carbon support films at 75KV, but have seen nice images of CNTs at 200kV. Any suggestions?

Also, do CNTs generate enough contrast on their own, or could negative staining be used to help improve the fine surface detail required to observe or amplify the subtle defects I am supposed to see. Thanks in advance for any suggestions, before I begin imaging.

Thanks, Mark Grimson
Texas Tech University

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Email: jtaillon-at-umd.edu
Name: Josh Taillon

Organization: University of Maryland

Title-Subject: [Filtered] Reading metadata from FEI TIFF files

Message: Hello,

I am wondering if anyone knows of a program that can extract the
metadata from TIFF files saved by an FEI Helios Nanolab 650. There is an
incredible amount of data that can be read by right-clicking into
Properties on the TIFF file (when using the FEI support PC), but I
cannot seem to find a program that can read this data offline. I figure
it might be somehow proprietary, but thought I would email the list in
hopes that someone has some experience with this.

Thank you,
Josh Taillon

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Email: behl-at-csulb.edu
Name: Richard Behl

Organization: California State University Long Beach

Title-Subject: [Filtered] Oxford EDS detector window

Message: Hi-
We have an Oxford Inca Energy EDS system purchased in 2002 with a "X-ray
detector: Si(Li) 138eV or better guaranteed, 10mm2, with SATW (Super
Atmospheric Thin Window) for detection of B - U" on the invoice and
quote. On the actual detector, the window is labeled "ATW2"

Can anyone tell me the material that this window is made of and its
thickness?

Thank you!

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Hello Josh,
try to use Graphicconverter, if you work MAC-based. If it cannot read the metadata you can ask Thorsten Lemke at
http://www.lemkesoft.com/ to do a special add-on for you to read out the data into an EXCEL sheet or something like that.

...Just a satisfied customer.

Best wishes,
Stefan

-----------------------------------------------------
Stefan Diller - Scientific Photography
Arndtstrasse 22
D - 97072 Wuerzburg Germany
++49-931-7848700 Phone
++49-931-7848701 Fax
++49-175-7177051 Mobile

Websites:
www.stefan-diller.com
www.electronmicroscopy.info
www.elektronenmikroskopie.info
www.assisi.de
www.zwillingsprojekt.de
Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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} Title-Subject: [Filtered] Reading metadata from FEI TIFF files
}
} Message: Hello,
}
} I am wondering if anyone knows of a program that can extract the
} metadata from TIFF files saved by an FEI Helios Nanolab 650. There is an
} incredible amount of data that can be read by right-clicking into
} Properties on the TIFF file (when using the FEI support PC), but I
} cannot seem to find a program that can read this data offline. I figure
} it might be somehow proprietary, but thought I would email the list in
} hopes that someone has some experience with this.
}
} Thank you,
} Josh Taillon
}
}
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10, 23 -- From stefan.diller-at-t-online.de Sat Nov 3 02:41:42 2012
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Hello,

I usually open the FEI tiff file using the Notepad or other text viewer. You will see at the start a lot of gibberish, but going to the end of the file you will see all the metadata. You will need to figure out what they mean, but most of them are pretty straightforward.

Hope this helps,

Carlos

—
Carlos Kazuo Inoki

On Nov 3, 2012, at 4:58 AM, "microscopylistserver-noreply-at-microscopy.com" {microscopylistserver-noreply-at-microscopy.com} wrote:

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} Title-Subject: [Filtered] Reading metadata from FEI TIFF files
}
} Message: Hello,
}
} I am wondering if anyone knows of a program that can extract the
} metadata from TIFF files saved by an FEI Helios Nanolab 650. There is an
} incredible amount of data that can be read by right-clicking into
} Â“PropertiesÂ” on the TIFF file (when using the FEI support PC), but I
} cannot seem to find a program that can read this data offline. I figure
} it might be somehow proprietary, but thought I would email the list in
} hopes that someone has some experience with this.
}
} Thank you,
} Josh Taillon
}
}
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8, 28 -- From carlos.inoki-at-lnls.br Sat Nov 3 10:17:21 2012
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Our Bio-formats projects is designed to open�proprietary�microscopy formats. This library can be used in many programs including ImageJ, seehttp://loci.wisc.edu/software/bio-formats

We currently support 127 formats and are trying to add support for electron microscopy formats include FEI tiff. See:
http://loci.wisc.edu/bio-formats-format/fei-tiff

best
kevin

On 11/03/12, carlos.inoki-at-lnls.br wrote:
}
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} Hello,
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} I usually open the FEI tiff file using the Notepad or other text viewer. You will see at the start a lot of gibberish, but going to the end of the file you will see all the metadata. You will need to figure out what they mean, but most of them are pretty straightforward.
}
} Hope this helps,
}
} Carlos
}
} —
} Carlos Kazuo Inoki
}
} On Nov 3, 2012, at 4:58 AM, "microscopylistserver-noreply-at-microscopy.com" {microscopylistserver-noreply-at-microscopy.com} wrote:
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} } Title-Subject: [Filtered] Reading metadata from FEI TIFF files
} }
} } Message: Hello,
} }
} } I am wondering if anyone knows of a program that can extract the
} } metadata from TIFF files saved by an FEI Helios Nanolab 650. There is an
} } incredible amount of data that can be read by right-clicking into
} } Â“PropertiesÂ” on the TIFF file (when using the FEI support PC), but I
} } cannot seem to find a program that can read this data offline. I figure
} } it might be somehow proprietary, but thought I would email the list in
} } hopes that someone has some experience with this.
} }
} } Thank you,
} } Josh Taillon
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} 8, 28 -- From carlos.inoki-at-lnls.br Sat Nov 3 10:17:21 2012
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--
Kevin W. Eliceiri
Director,�Laboratory for Optical and Computational Instrumentation (LOCI)
Departments Cell and Molecular Biology and Biomedical Engineering
Affiliate Principal Investigator, Morgridge Institute for Research (MIR)
Room 271 Animal Sciences,�1675 Observatory Drive,�Madison, WI 53706
Phone: 608-263-6288

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Josh,

Anyone can also read the metadata from most tiff files using any simple text reader such as Notepad. Open the file with notepad and scroll down to the bottom of the file. You will see metadata similar to the following:

[Beam]
HV=20000
Spot=5
StigmatorX=0.000550687
StigmatorY=-0.000959411
BeamShiftX=0
BeamShiftY=0
ScanRotation=0
ImageMode=Normal
Beam=EBeam
Scan=EScan

Joe Neilly
Abbott Laboratories
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Title-Subject: [Filtered] Reading metadata from FEI TIFF files

Message: Hello,

I am wondering if anyone knows of a program that can extract the
metadata from TIFF files saved by an FEI Helios Nanolab 650. There is an
incredible amount of data that can be read by right-clicking into
�"Properties�" on the TIFF file (when using the FEI support PC), but I
cannot seem to find a program that can read this data offline. I figure
it might be somehow proprietary, but thought I would email the list in
hopes that someone has some experience with this.

Thank you,
Josh Taillon

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I recently tilted a very thin foil of aluminum to the {110} zone axis, carefully aligned the instrument, inserted a large objective aperture, and attempted lattice imaging of fine precipitates. Much to my surprise, I obtained lattice images of the Al {110} zone. A Fourier transform of the HR lattice image, using my image-space magnification calibration, shows spacings of 0.2335 and 0.2034 nm in a clear {110} -shaped diffractogram. Looking up the lattice spacings (UIUC WebEMAPS website), I find values of 0.2338 and 0.2025 nm for the (111) and (002) planes in aluminum, so this makes me think I'm seeing a true Al {110} lattice image.

However, I'm using a CM200FEG with SuperTwin lens, which should give Cs=1.2 mm and Cc=1.2 mm. Using the equation for point resolution in De Graef's book (P. 601), and plugging in Cs=1.2e-3 m and lambda = 2.508e-12 m, I calculate ps~0.237 nm, which makes me think I shouldn't be able to see these planes -- and that neglects Cc and delta-E, so resolution ought to be even worse.

Am I misunderstanding point resolution, and need to calculate a different (more forgiving) resolution function? Or am I imaging some sort of artifact (surface oxide?) that is convincingly similar to my real information?

Thanks,
Chad Parish

==============================Original Headers==============================
6, 30 -- From parishcm-at-ornl.gov Mon Nov 5 08:40:05 2012
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Hi Chad,

Since you are using a FEG source, the contrast transfer function (CTF)
doesn't damp very quickly. Normally point resolution is quoted as the
Scherzer resolution i.e. at the first zero of the CTF at a particular
defocus. In LaB6 TEMs, the source adds a damping function to the CTF so
you usually don't see much beyond that 1st zero. With a FEG, you have
much less damping of the CTF and the oscillations of the CTF start to
affect your image. Remember that the CTF indicates information transfer
through the scope, so if the CTF is non-zero you are passing
information. If the particular spatial frequencies are between the 1st
and 2nd zeros of the the CTF, you will see those frequencies in your image.

We've seen 1.5A information on a 200kV SuperTwin FEG by adjusting the
objective defocus so that the CTF passes those frequencies.
Interpretation should be done very carefully on 2nd and 3rd zone images.

Cheers,
Henk

At 11/5/2012 9:40 AM, parishcm-at-ornl.gov wrote:
}
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} I recently tilted a very thin foil of aluminum to the {110} zone axis, carefully aligned the instrument, inserted a large objective aperture, and attempted lattice imaging of fine precipitates. Much to my surprise, I obtained lattice images of the Al {110} zone. A Fourier transform of the HR lattice image, using my image-space magnification calibration, shows spacings of 0.2335 and 0.2034 nm in a clear {110} -shaped diffractogram. Looking up the lattice spacings (UIUC WebEMAPS website), I find values of 0.2338 and 0.2025 nm for the (111) and (002) planes in aluminum, so this makes me think I'm seeing a true Al {110} lattice image.
}
} However, I'm using a CM200FEG with SuperTwin lens, which should give Cs=1.2 mm and Cc=1.2 mm. Using the equation for point resolution in De Graef's book (P. 601), and plugging in Cs=1.2e-3 m and lambda = 2.508e-12 m, I calculate ps~0.237 nm, which makes me think I shouldn't be able to see these planes -- and that neglects Cc and delta-E, so resolution ought to be even worse.
}
} Am I misunderstanding point resolution, and need to calculate a different (more forgiving) resolution function? Or am I imaging some sort of artifact (surface oxide?) that is convincingly similar to my real information?
}
} Thanks,
} Chad Parish
}
}
}
} ==============================Original Headers==============================
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

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Hi all,
�
I am writing a list�of equipment for EM�and have been asked to find an approximate price in the market for used equipment.
I would be grateful if you could give me the (approximate) price of the listed equipment new or used but in perfect condition (the equipment was never used):
�
- Plate Degasser PD3 from Edwards (+ accessories to couple to the Edwards pump)
- Rotary pump RV3 from Edwards
- Anti-static device from Leica "Static line EN SL II" with power generator
- Leica cryochamber EM FC6 with isolated tube, the Dewar adapter, the Dewar itself (there is no indication on it so I don't know its capacity or even its name) and a cryotrim 45 knife.
�
All are in perfect condition, they have been�protected from dust, temperature changes and shock.
�
You may use dollars or euros.
�
Please note that I am only responsible for preparing the list, I am not responsible for selling the equipment so don't contact me to tell me that you are interested.
I will post a new message on the Listserver if/when we come to the point of selling them.
�
Many thanks in advance
�
Stephane

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NCSM/MSA members and microscopist in northern California,

Greetings and welcome to our call for your attendance at the Fall 2012
meeting, hosted by the Diabetes Center Microscopy Core, UCSF, and Larry
Ackerman. The venue is now finalized and we are all in for a good
evening of socializing, business discussions and scientific
presentations.

The meeting will start at 6:00 on Thursday, Nov 15th in the lobby
outside of room N114, Genentech Hall on the Mission Bay campus of UCSF.
A map of the campus, including parking and public transportation stops,
is in the URL, below.

6 - 6:30: Registration and socializing
6:30: Dinner, as we move into the conference room
7:00 - 7:30: Business Meeting (NCSM President)
7:30 - 9:00: Scientific Talks:
-Anna Celli, PhD., Department of Dermatology, UCSF: "FLIM (Fluorescence
Lifetime Imaging Microscopy) in Dermatology Research"
-Luis Comolli, PhD., Biophysics, UC Berkeley: Structural Analysis of
Lysinibacillus sphaericus S-layer by 2D and 3D Cryo-TEM.
-Tom Goddard, Chimera Developer, UCSF: Chimera Intro for Microscopy.
-9:00 Wrap-up, desert, socializing, networking

To finalize preparations for the meeting we must know your intentions
for attending so please contact me ASAP. And note that all attendees
will have to sign in with security in the lobby of Genentech Hall.

See you in two weeks.

Steve Samuelsson
President, NCSM

Mission Bay map:
http://www.ucsf.edu/sites/default/files/documents/ucsf_mission_bay_web.pdf,
also attached to this email; parking structures are clearly indicated.

--
Steven J. Samuelsson, Ph.D.
Principal Scientist
Director, Cell and Molecular Imaging
Biosciences Division
SRI International, Inc.
333 Ravenswood Ave.
Menlo Park, CA 94025
(650) 859-2980 office
(650) 859-3153 fax

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Do you have a young person on your holiday gift list? Are you
considering a 'first microscope' plus a book about the microworld? If
so, MSA's Project MICRO can help you make a selection; see the URL
below. MICRO's 'buying microscopes' page gives advice on selecting
the right scope, and the booklist (over 250 reviewed entries!) has a
detailed, efficient search engine that will help you find the best
book. If the website doesn't meet your needs, please Email me
directly.
--
Caroline Schooley
Project MICRO Coordinator
Microscopy Society of America
45301 Caspar Point Road, Box 117
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While it is possible to use a text editor (Notepad, Notepad++, TextPad) to find TIFF metadata, the flexibility of the TIFF format doesn't always make it easy. The metadata can be stored anywhere, the beginning, the end or the middle of the file. However tools like http://meta-extractor.sourceforge.net/documentation.htm or http://www.awaresystems.be/imaging/tiff/astifftagviewer.html can reliably extract metadata from TIFF files either using a friendly GUI or an automatable command line interface. Usually the meta data you want will be associated with the "ImageDescription" tag (code 270 or 0x010E) although there are numerous other tags that could contain useful information. (see http://www.awaresystems.be/imaging/tiff/tifftags/baseline.html) Some vendors even implement custom tags and register these with Adobe for special data types like x-ray spectrum data. (see http://partners.adobe.com/public/developer/tiff/index_reg.html)

Nicholas

=======================================
Nicholas W. M. Ritchie
Physicist, Surface and Microanalysis Science Division
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100 Bureau Drive, MS: 8371
Gaithersburg, MD 20899-8371
(Work) (301) 975-3929 (Cell) (240) 883-8982

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Title-Subject: [Filtered] Reading metadata from FEI TIFF files

Message: Hello,

I am wondering if anyone knows of a program that can extract the metadata from TIFF files saved by an FEI Helios Nanolab 650. There is an incredible amount of data that can be read by right-clicking into �"Properties�" on the TIFF file (when using the FEI support PC), but I cannot seem to find a program that can read this data offline. I figure it might be somehow proprietary, but thought I would email the list in hopes that someone has some experience with this.

Thank you,
Josh Taillon

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Chad;
You might get some insights by downloading Max Sidorov's "CTF Explore= r" at:

http://www.maxsidorov.com/ctfexplorer/

You will be able to see how resolution beyond the point resolution can be achieved with a FEG. The only catch in real microscopy is that the waves carrying resolution beyond the point limit may be out of phase with waves carrying resolution at different frequencies, and the resulting images may not be directly interpretable representations of the structure of you specimen. Also if you study John Spence's classic text on HRTEM you will have a good grasp of what is going on here.

John Mardinly
ASU

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I recently tilted a very thin foil of aluminum to the {110} zone axis, carefully aligned the instrument, inserted a large objective aperture, and attempted lattice imaging of fine precipitates. Much to my surprise, I obtained lattice images of the Al {110} zone. A Fourier transform of the HR lattice image, using my image-space magnification calibration, shows spacings of 0.2335 and 0.2034 nm in a clear {110} -shaped diffractogram. Looking up the lattice spacings (UIUC WebEMAPS website), I find values of 0.2338 and 0.2025 nm for the (111) and (002) planes in aluminum, so this makes me think I'm seeing a true Al {110} lattice image.

However, I'm using a CM200FEG with SuperTwin lens, which should give Cs=1.2 mm and Cc=1.2 mm. Using the equation for point resolution in De Graef's book (P. 601), and plugging in Cs=1.2e-3 m and lambda = 2.508e-12 m, I calculate ps~0.237 nm, which makes me think I shouldn't be able to see these planes -- and that neglects Cc and delta-E, so resolution ought to be even worse.

Am I misunderstanding point resolution, and need to calculate a different (more forgiving) resolution function? Or am I imaging some sort of artifact (surface oxide?) that is convincingly similar to my real information?

Thanks,
Chad Parish

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Title-Subject: [Filtered] Reading metadata from FEI TIFF files

Message: Hello everyone,

Thank you for the wide range of responses. I just wanted to give a
summary of what I have learned so that it may be useful for other people.

The FEI metadata fields are definitely accessible just by opening the
TIFF file as a text file in your favorite text editor. This is more than
sufficient if all you're doing is checking a number quickly (Thank you
to everyone that pointed this out to me).

Using the AsTiffTagViewer
(http://www.awaresystems.be/imaging/tiff/astifftagviewer.html) (Thank
you, Nicholas Ritchie), I was able to find that the codes are stored as
ASCII text in tag number 34682 (for the FEI Helios, at least). This is a
non-standard tag, but is freely readable using Tiff libraries.

The most useful website I found was
http://www.farsight-toolkit.org/wiki/FARSIGHT_Tutorials/Bio-Formats
(Thank you Kevin, from Wisconsin for pointing me in the right
direction). This is a simple tutorial that will lead you through
installing the Bio-formats command line tools. They have developed a
plugin to read specifically from FEI Tiffs, and the `showinf` command
will cleanly display all the custom FEI tags that are present in the
file. Using this, you can then `grep` or search for whatever particular
information you need. I think this will be the solution I use going forward.

Thank you everyone for all of your help. I certainly learned a lot.

- Josh

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Greetings

I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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Jon,

Diatomaceous earth would be good. Soft, sedimentary rock made of
diatoms, used for filters, mild abrasives (some toothpastes, although
I can't think of a brand off the top of my head), and the like.
Also, have a chat with the geologists in your area - lots of the
limestones in California are composed mostly of forams, ostracods,
and suchlike critters.
Along with dragging a plankton net through SF bay ...

Phil

} Greetings
}
} I would really like to get my hands on some micro biological samples
} suitable for SEM. The kinds of things I am thinking about are
} radiolarians, forams, coccoliths, diatoms, that kind of thing. Any
} ideas about sources of raw materials I can get to provide beginning
} SEM students with a surprise at the microscope?
}
} Thanks
}
} Jon
}
} Jonathan Krupp
} Delta College
} 5151 Pacific Ave.
} Box 212
} Stockton, CA 95207
} 209-954-5284
} jkrupp-at-deltacollege.edu
}
} Find us on Facebook -at-
} Electron Microscopy at SJ Delta College

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Jon,

I have a few environmental scientists from down the hall who I image diatoms for on our SEM. I believe they have small glass diatom "traps" they use to collect samples, though I'm fairly sure you can pick up almost any stone in a healthy stream and scrape some off.

As far as sample prep goes, they acidify the diatoms to eliminate biological material, then drip a slurry onto an Al stub and dry the sample. It seems to work pretty well and I've been able to get great images of a number of different genera.

Cheers,

Rob Dean
Department of Earth Sciences
Dickinson College
�
* Please consider your environmental responsibility before printing this e-mail or any other document.

-----Original Message-----
X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
Sent: Wednesday, November 07, 2012 3:50 PM
To: Dean, Robert

Greetings

I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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Hi Jon & Rob,

If you are looking for diatoms, why not run over to your local home
improvement or swimming pool supplier and pick up some diatomaceous
earth. It is often used as filtering material for swimming pools and
there are now some eco friendly insecticides that use diatomaceous
earth. (do a net search)

You will have to suspend the material to separate the diatoms from the
rest of the "earth", but it is a cheap source of diatoms!

Cheers,
Henk

At 11/7/2012 4:28 PM, deanr-at-dickinson.edu wrote:
}
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} Jon,
}
} I have a few environmental scientists from down the hall who I image diatoms for on our SEM. I believe they have small glass diatom "traps" they use to collect samples, though I'm fairly sure you can pick up almost any stone in a healthy stream and scrape some off.
}
} As far as sample prep goes, they acidify the diatoms to eliminate biological material, then drip a slurry onto an Al stub and dry the sample. It seems to work pretty well and I've been able to get great images of a number of different genera.
}
} Cheers,
}
} Rob Dean
} Department of Earth Sciences
} Dickinson College
}
} * Please consider your environmental responsibility before printing this e-mail or any other document.
}
}
} -----Original Message-----
} X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
} Sent: Wednesday, November 07, 2012 3:50 PM
} To: Dean, Robert
} Subject: [Microscopy] Micro SEM samples
}
}
}
}
} ----------------------------------------------------------------------------
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} ----------------------------------------------------------------------------
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} Greetings
}
} I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?
}
} Thanks
}
} Jon
}
} Jonathan Krupp
} Delta College
} 5151 Pacific Ave.
} Box 212
} Stockton, CA 95207
} 209-954-5284
} jkrupp-at-deltacollege.edu
}
} Find us on Facebook -at-
} Electron Microscopy at SJ Delta College
}
}
}
}
}
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--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

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Contents Retrieved from Microscopy Listserver Archives
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Hi Jon & Rob,

If you are looking for diatoms, why not run over to your local home
improvement or swimming pool supplier and pick up some diatomaceous
earth. It is often used as filtering material for swimming pools and
there are now some eco friendly insecticides that use diatomaceous
earth. (do a net search)

You will have to suspend the material to separate the diatoms from the
rest of the "earth", but it is a cheap source of diatoms!

Cheers,
Henk

At 11/7/2012 4:28 PM, deanr-at-dickinson.edu wrote:
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Jon,
}
} I have a few environmental scientists from down the hall who I image diatoms for on our SEM. I believe they have small glass diatom "traps" they use to collect samples, though I'm fairly sure you can pick up almost any stone in a healthy stream and scrape some off.
}
} As far as sample prep goes, they acidify the diatoms to eliminate biological material, then drip a slurry onto an Al stub and dry the sample. It seems to work pretty well and I've been able to get great images of a number of different genera.
}
} Cheers,
}
} Rob Dean
} Department of Earth Sciences
} Dickinson College
}
} * Please consider your environmental responsibility before printing this e-mail or any other document.
}
}
} -----Original Message-----
} X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
} Sent: Wednesday, November 07, 2012 3:50 PM
} To: Dean, Robert
} Subject: [Microscopy] Micro SEM samples
}
}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Greetings
}
} I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?
}
} Thanks
}
} Jon
}
} Jonathan Krupp
} Delta College
} 5151 Pacific Ave.
} Box 212
} Stockton, CA 95207
} 209-954-5284
} jkrupp-at-deltacollege.edu
}
} Find us on Facebook -at-
} Electron Microscopy at SJ Delta College
}
}
}
}
}
}
}
}
} ==============================Original Headers==============================
} 13, 30 -- From jkrupp-at-deltacollege.edu Wed Nov 7 14:42:02 2012
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} 25, 23 -- From deanr-at-dickinson.edu Wed Nov 7 15:27:23 2012
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} 25, 23 -- To: "'Microscopy-at-microscopy.com' (Microscopy-at-microscopy.com)"
} 25, 23 -- {Microscopy-at-microscopy.com}
} 25, 23 -- Subject: [Microscopy] RE: Micro SEM samples
} 25, 23 -- Thread-Topic: [Microscopy] RE: Micro SEM samples
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}

--
Hendrik O. Colijn www.ceof.ohio-state.edu
OSU Campus Electron Optics Facility colijn.1-at-osu.edu
040 Fontana Labs (614) 292-0674 (V)
116 W. 19th Ave. (614) 292-7523 (F)
Columbus, OH 43210

"Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."

==============================Original Headers==============================
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11, 40 -- From: Henk Colijn {colijn.1-at-osu.edu}
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11, 40 -- {microscopy-at-microscopy.com}
11, 40 -- Subject: Re: [Microscopy] RE: Micro SEM samples
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Contents Retrieved from Microscopy Listserver Archives
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I did just that on several occasions. Ji was able to go to a local
pool supply place and just sweep about a square foot of the floor near
the diatomaceous earth stack and got enough sample to make several
stubs.

--Justin.

"America believes in education; the average professor earns more money
in a year than a professional athlete earns in a whole week." Evan
Esar

On Nov 7, 2012, at 5:15 PM, "colijn.1-at-osu.edu" {colijn.1-at-osu.edu} wrote:

}
}
}
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} ----------------------------------------------------------------------------
}
} Hi Jon & Rob,
}
} If you are looking for diatoms, why not run over to your local home
} improvement or swimming pool supplier and pick up some diatomaceous
} earth. It is often used as filtering material for swimming pools and
} there are now some eco friendly insecticides that use diatomaceous
} earth. (do a net search)
}
} You will have to suspend the material to separate the diatoms from the
} rest of the "earth", but it is a cheap source of diatoms!
}
} Cheers,
} Henk
}
}
}
} At 11/7/2012 4:28 PM, deanr-at-dickinson.edu wrote:
} }
} }
} } ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------------
} }
} } Jon,
} }
} } I have a few environmental scientists from down the hall who I image diatoms for on our SEM. I believe they have small glass diatom "traps" they use to collect samples, though I'm fairly sure you can pick up almost any stone in a healthy stream and scrape some off.
} }
} } As far as sample prep goes, they acidify the diatoms to eliminate biological material, then drip a slurry onto an Al stub and dry the sample. It seems to work pretty well and I've been able to get great images of a number of different genera.
} }
} } Cheers,
} }
} } Rob Dean
} } Department of Earth Sciences
} } Dickinson College
} }
} } * Please consider your environmental responsibility before printing this e-mail or any other document.
} }
} }
} } -----Original Message-----
} } X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
} } Sent: Wednesday, November 07, 2012 3:50 PM
} } To: Dean, Robert
} } Subject: [Microscopy] Micro SEM samples
} }
} }
} }
} }
} } ----------------------------------------------------------------------------
} } The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
} } To Subscribe/Unsubscribe -- http://www.microscopy.com/MicroscopyListserver
} } On-Line Help http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------------
} }
} } Greetings
} }
} } I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?
} }
} } Thanks
} }
} } Jon
} }
} } Jonathan Krupp
} } Delta College
} } 5151 Pacific Ave.
} } Box 212
} } Stockton, CA 95207
} } 209-954-5284
} } jkrupp-at-deltacollege.edu
} }
} } Find us on Facebook -at-
} } Electron Microscopy at SJ Delta College
} }
} }
} }
} }
} }
} }
} }
} }
} } ==============================Original Headers==============================
} } 13, 30 -- From jkrupp-at-deltacollege.edu Wed Nov 7 14:42:02 2012
} } 13, 30 -- Received: from mailin.deltacollege.edu (mailin.deltacollege.edu [207.62.178.150])
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} } 13, 30 -- Content-Type: text/plain; charset=us-ascii
} } 13, 30 -- Subject: Micro SEM samples
} } 13, 30 -- Date: Wed, 7 Nov 2012 12:42:01 -0800
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} }
} } ==============================Original Headers==============================
} } 25, 23 -- From deanr-at-dickinson.edu Wed Nov 7 15:27:23 2012
} } 25, 23 -- Received: from exmail.dickinson.edu (exmail2.DICKINSON.EDU [64.9.56.29])
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} } 25, 23 -- ([::1]) with mapi id 14.01.0355.002; Wed, 7 Nov 2012 16:27:22 -0500
} } 25, 23 -- From: "Dean, Robert" {deanr-at-dickinson.edu}
} } 25, 23 -- To: "'Microscopy-at-microscopy.com' (Microscopy-at-microscopy.com)"
} } 25, 23 -- {Microscopy-at-microscopy.com}
} } 25, 23 -- Subject: [Microscopy] RE: Micro SEM samples
} } 25, 23 -- Thread-Topic: [Microscopy] RE: Micro SEM samples
} } 25, 23 -- Thread-Index: Ac29LgGwY06g7eikRk+dMRjICnDmnw==
} } 25, 23 -- Date: Wed, 7 Nov 2012 21:27:21 +0000
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}
} --
} Hendrik O. Colijn www.ceof.ohio-state.edu
} OSU Campus Electron Optics Facility colijn.1-at-osu.edu
} 040 Fontana Labs (614) 292-0674 (V)
} 116 W. 19th Ave. (614) 292-7523 (F)
} Columbus, OH 43210
}
} "Time is that quality of nature which keeps things from happening all at once. Lately it doesn't seem to be working."
}
}
}
} ==============================Original Headers==============================
} 11, 40 -- From colijn.1-at-osu.edu Wed Nov 7 16:09:37 2012
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Jon,
Check out this website so the students can compare their findings. http://www.ucl.ac.uk/GeolSci/micropal/diatom.html

Lita Duraine
EM Technologist
Bellen Lab
HHMI- Molecular Genetics
Duncan Neurological Research Institute
1250 Moursund St.
Houston, TX 77030
Rm: N1165.17
MS: N1125.50
832-824-8772 TEM Room
979-549-6526 Cell
http://flypush.imgen.bcm.tmc.edu/lab/people/lita.php

-----Original Message-----
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Sent: Wednesday, November 07, 2012 2:55 PM
To: Duraine, Lita

Greetings

I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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23, 28 -- From duraine-at-bcm.edu Wed Nov 7 18:21:45 2012
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Hi Folks,

I have an epi illumination setup on an old Leitz Metalloplan with DIC, and I am seeing double images as soon as I insert the Nomarski prism. The two images overlay something like a few microns at ~200x. When I change objectives, the shift stays constant (in angle), while the objects in the view change size. So, at 200x, the shift may be 2 microns, but at 400x the shift is now 1 micron. I can adjust the polarizers so the double image is not visible, but then I don't get any extra contrast. I've tried adjusting the optical column, and the shift is insensitive to the incidence angle or focus of the illumination system. Moving the Nomarski prism up and down several mm, yaw or pitch several degrees makes no difference in the double image, though it will eventually leave me with a fuzzy image.

Is this just an incorrect Nomarski prism for low mag? I notice at highest mag (about 80x objective and 18x eyepiece), the shift is just slightly more than the point resolution so it slightly degrades the quality of the image, though the contrast increases. So I hypothesize that the prism was chosen to optimize DIC contrast for the highest mag (and even not quite right for that), leaving the lower mags with a double image. However, if I am doing something wrong, I'd like to fix it.

Any ideas?

Cheers,

Zack Gainsforth
Space Sciences Laboratory, UC Berkeley
7 Gauss Way
Berkeley, CA 94720
510-642-9733
zackg-at-ssl.berkeley.edu

==============================Original Headers==============================
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And there's also kitty litter - I think this is still mostly diatomaceous
earth.
cheers,
Rosemary

Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
M 61 2 420 972 028
E rosemary.white-at-csiro.au

On 8/11/12 9:15 AM, "colijn.1-at-osu.edu" {colijn.1-at-osu.edu} wrote:

}
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Ha,

I can. I used them at a previous position.
Run. As fast as you can. NOT a good idea!

Phil

} Email: mekeel-at-med.unc.edu
} Name: Ha Mekeel
}
} Organization: University of North Carolina
}
} Title-Subject: [Filtered] Specialty Underwriters
}
} Message: Can anyone give me an evaluation of an organization called
} Specialty Underwriters? We have
} a Tecnai 12 and were stunned to find that the SU was contracted
} without including the microscopists
} in the decision. What experiences have you had.

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Many thanks to the people who replied, both on- and off-list, to my query.

The answer appears to be, if I am understanding and summarizing correctly, that point resolution (~0.24 nm for a CM200FEG-ST) is a less appropriate measure than information limit (~0.14 nm). The FEG source results in a slower damping of the transfer function, so although the Al lattice spacings are past the first zero of the function, the transfer of information is generally not zero at those spacings, so the information makes it to the camera. Several people cautioned me that since we're beyond the first pass band I should exercise extreme caution in interpretation of the image information. However, since the interest of the experiment was to see if precipitates aligned along given lattice directions, and was not structure determination, I think I am safe in that regard.

Again, thanks to everyone who replied!

Chad

PS: I was able to convince myself with the following simple MATLAB script tested under Version 7.10.0.499 (R2010a) --

clear all; close all; pack; memory
chi = -at-(u,df,l,Cs) pi.*df.*l.*u.*u+0.5.*pi.*Cs.*l.*l.*l.*u.*u.*u.*u; %transfer function
l=2.508e-3; %wavelength in nm
Cs=1.2e6; % Cs in nm
u=0.01:0.01:6; % spacing in inverse nm
dfSCH=-1.2*(Cs*l)^0.5; % Scherzer defocus
plot(u,sin(chi(u,dfSCH,l,Cs)))
hold all
u111=1/.2338;
u200=1/.2025;
plot(u111,sin(chi(u111,dfSCH,l,Cs)),'o')
plot(u200,sin(chi(u200,dfSCH,l,Cs)),'*')

-----Original Message-----
X-from: parishcm-at-ornl.gov [mailto:parishcm-at-ornl.gov]
Sent: Monday, November 05, 2012 9:47 AM
To: Parish, Chad M.

I recently tilted a very thin foil of aluminum to the {110} zone axis, carefully aligned the instrument, inserted a large objective aperture, and attempted lattice imaging of fine precipitates. Much to my surprise, I obtained lattice images of the Al {110} zone. A Fourier transform of the HR lattice image, using my image-space magnification calibration, shows spacings of 0.2335 and 0.2034 nm in a clear {110} -shaped diffractogram. Looking up the lattice spacings (UIUC WebEMAPS website), I find values of 0.2338 and 0.2025 nm for the (111) and (002) planes in aluminum, so this makes me think I'm seeing a true Al {110} lattice image.

However, I'm using a CM200FEG with SuperTwin lens, which should give Cs=1.2 mm and Cc=1.2 mm. Using the equation for point resolution in De Graef's book (P. 601), and plugging in Cs=1.2e-3 m and lambda = 2.508e-12 m, I calculate ps~0.237 nm, which makes me think I shouldn't be able to see these planes -- and that neglects Cc and delta-E, so resolution ought to be even worse.

Am I misunderstanding point resolution, and need to calculate a different (more forgiving) resolution function? Or am I imaging some sort of artifact (surface oxide?) that is convincingly similar to my real information?

Thanks,
Chad Parish

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I suggest pollen. Most of the samples I'm familiar with dry nicely, need a gold coat for conductivity and look great. Different hair with their scale patterns is always interesting. You can make scale cast and see it with the light microscope and then show the same hair in the SEM. Try dog, cat human and wool fibers. Cross sections of wood can be interesting. They can take a little more care in prepping the samples. I'd put bamboo, a grass, in your wood selection just for contrast. Don't forget balsa wood and cork.

Have fun and let us know what you use.

Frank

-----Original Message-----
X-from: jkrupp-at-deltacollege.edu [mailto:jkrupp-at-deltacollege.edu]
Sent: Wednesday, November 07, 2012 3:54 PM
To: Frank Karl

Greetings

I would really like to get my hands on some micro biological samples suitable for SEM. The kinds of things I am thinking about are radiolarians, forams, coccoliths, diatoms, that kind of thing. Any ideas about sources of raw materials I can get to provide beginning SEM students with a surprise at the microscope?

Thanks

Jon

Jonathan Krupp
Delta College
5151 Pacific Ave.
Box 212
Stockton, CA 95207
209-954-5284
jkrupp-at-deltacollege.edu

Find us on Facebook -at-
Electron Microscopy at SJ Delta College

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X-from: Richard Wuhrer {richard.wuhrer-at-uws.edu.au}

Dear All,

The 12th Biennial Australian Microbeam Analysis Society Symposium (AMAS XII) will be held in at the
University of Technology, Sydney from the 4th to 8th February 2013.

http://www.microscopy.org.au/amas/amas12/

The aim of the AMAS Symposium is to provide a forum where participants can discuss and share ideas
on current advances, trends and challenges in microanalysis and imaging, with an emphasis on
practical solutions and applications. We strongly encourage you to present your research work using
scanning or transmission electron microscopy and microanalysis.

A wide range of introductory and advanced workshops in the general area of microscopy and
microanalysis will be run prior to the Symposium on the 4th and 5th February 2013.

Visit http://www.microscopy.org.au/amas/amas12/ for further information on; registration,
accommodation, call for papers, student travel bursary information as well as the technical and
social program.

Could you please distribute this email to your colleagues and anyone that you think might be
interested in attending.

Best regards,

Richard Wuhrer (UWS) Co-Chair
richard.wuhrer-at-uws.edu.au

**

*Dr Richard Wuhrer*
Research Manager
Advanced Materials Characterisation Facility
University of Western Sydney
Office of Pro-Vice Chancellor (Research)
Academic and Research Division
Locked Bag 1797 Penrith NSW 2751 Australia
*Mobile: 0411 877 476*
Phone: +61 2 9685 9089
Fax: +61 2 9685 9915
Email:*Richard.Wuhrer-at-uws.edu.au {mailto:Richard.Wuhrer-at-uws.edu.au} *
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I also saw the Dino-Lite demonstrated at a trade show, and decided to
purchase. We use it for photographing Hallmarks, and other identification
data on jewelry items prior to metallographic analysis. The built in LEDs do
not always provide the best illumination. For our purposes, we often use
photo floods on a copy stand (actually from an MP-4) with a white paper
diffuser around the sample. The Dino-Lite stand is a must. Hand-holding the
device still at several x magnification is impossible. The software hangs
every once in a while, put we are generally happy with the product. Our real
workhorse for macro work is a Canon digital camera with ultra close macro.

Jeff Stewart
Metallographic Laboratory Manager
LeachGarner
49 Pearl Street
Attleboro, MA 02703
508-222-7400 x1329
----- Original Message -----
X-from: {werner1-at-slb.com}
To: {jeff-at-metallography.com}
Sent: Wednesday, October 20, 2010 1:51 PM

Hello,

We are evaluating to purchase a dual-beam FIB for our lab. It is going to be used mainly for TEM sample preparation (semiconductor, ceramics, metal, soft...). Looking for the options available in the market, we narrow it down to FEI and Zeiss. Probably FEI Nanolab or Zeiss Auriga. The kind of information I am looking is about what is your experience with these instruments. Specially maintenance and how much if costs per year in real world (parts and service).

One advantage about Zeiss was the Argon beam for sample cleaning. But I was told that it is not available for the Auriga dual-beam and It is not really necessary to prepare good TEM samples. Tripod polishing produces wonderful samples, but most people have some difficulty in doing so. Did you stop using tripod polishing after start using FIB? Does low energy ion milling helps to improve the quality of FIBed samples (tripod quality)?

Any opinion or experience that you want to share will be very useful for us.

Thanks,

Carlos
____________________________________________
Carlos Kazuo Inoki
Brazilian Nanotechnology National Laboratory
Rua Giuseppe Maximo Scolfaro 10000
Campinas, SP 13083-970
Brazil

==============================Original Headers==============================
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Hello Carlos.

Just out of curiosity, is there any specific reason you're not looking
into the Tescan Lyra 3 dualbeam system?

Regards,
Roy

-----Original Message-----
X-from: carlos.inoki-at-lnls.br [mailto:carlos.inoki-at-lnls.br]
Sent: Saturday, November 10, 2012 00:56
To: Roy Golombick

Hello,

We are evaluating to purchase a dual-beam FIB for our lab. It is going
to be used mainly for TEM sample preparation (semiconductor, ceramics,
metal, soft...). Looking for the options available in the market, we
narrow it down to FEI and Zeiss. Probably FEI Nanolab or Zeiss Auriga.
The kind of information I am looking is about what is your experience
with these instruments. Specially maintenance and how much if costs per
year in real world (parts and service).

One advantage about Zeiss was the Argon beam for sample cleaning. But I
was told that it is not available for the Auriga dual-beam and It is not
really necessary to prepare good TEM samples. Tripod polishing produces
wonderful samples, but most people have some difficulty in doing so. Did
you stop using tripod polishing after start using FIB? Does low energy
ion milling helps to improve the quality of FIBed samples (tripod
quality)?

Any opinion or experience that you want to share will be very useful
for us.

Thanks,

Carlos
____________________________________________
Carlos Kazuo Inoki
Brazilian Nanotechnology National Laboratory Rua Giuseppe Maximo
Scolfaro 10000 Campinas, SP 13083-970 Brazil

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Analytical Instrumentation specialist position, Montclair State University

The Department of Earth and Environmental Studies at Montclair State University seeks an analytical instrumentation specialist to support research and teaching efforts. The specialist will assist students, faculty, and visitors in the use of our JY Horiba Ultima C
ICP-OES, Thermo X7 Series ICP-MS, and several other analytical instruments housed in the department (GC/MS, AA, XRD, HPLC, IC, TOC, CHNS analyzer, gamma detector) along with their supporting sample preparation equipment. The successful applicant will have expertise in the preparation and quantitative analysis of a wide range of geologic and environmental materials (rock, sediment, soil, water, plants), experience with maintenance and troubleshooting of one or more of the instruments listed above and their related ancillary equipment, and excellent communication and interpersonal skills with the ability to work in a multidisciplinary department with a wide variety of beginning through advanced users.

Primary responsibilities of the position include the training of new users on the analytical instruments and their supporting equipment, conducting basic preventative maintenance on the instruments, diagnosing instrument malfunctions, and coordinating with technical repair personnel from the manufacture, overseeing chemical hygiene and chemical safety in the analytical support labs for the instrumentation, and working with department chair on purchasing and billing.

A Masters degree in a STEM discipline and at least 3 years of experience in the operation and maintenance of ICP-OES/ICP-MS instruments and one or more of the instruments listed above and their sample preparation equipment is required. Strong communication skills and demonstrated experience in the preparation and analysis of a wide array of sample types is also required. Strong knowledge of computers, electronics, and troubleshooting is highly desirable. The successful candidate will be qualified to conduct consulting work, collaborative research with faculty/students, and/or pursue external research funding. Experience with other major analytical equipment is a plus.

Electronic applications, combined in a single .doc or .pdf file are preferred. Send to: Dr. Michael Kruge at krugem-at-mail.montclair.edu, Department of Earth and Environmental Studies, Montclair State University, Montclair, NJ 07043. Applications must include a CV, statement of technical skills, research and laboratory management experience, and the names and contact information for at least three references.

--
*********************
Stefanie A. Brachfeld
Associate Professor and Associate Chair
Geoscience Graduate Program Coordinator
Department of Earth & Environmental Studies
Montclair State University
Upper Montclair, NJ 07043
phone: (973) 655-5129
fax: 973-655-4072
brachfelds-at-mail.montclair.edu
*********************

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Does anyone have any experience with this? We have a Gatan CL system on a Hitachi 3400 and we were curious how beneficial a cooling system would be. Would a peltier stage be cold enough? Would an LN2 cold finger be too cold? Thanks.

ERiC Jay Miller
Microscopy & Imaging Specialist
Electron Probe Instrumentation Center

Northwestern University
Mail: 2036 Cook Hall
Office: 1152 Cook Hall
2220 Campus Drive
Evanston, IL 60208-3108

ph: (847) 467-0789
fax: (847) 467-657

http://www.nuance.northwestern.edu

==============================Original Headers==============================
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Eric,

We have the same system, and a Deben cold stage. Only goes down to
-30 C (high vacuum), but so far it's been all we need. Mostly for
mineralogical samples.
It will be interesting if anyone needs colder temperatures.

Phil

} Does anyone have any experience with this? We have a Gatan CL system
} on a Hitachi 3400 and we were curious how beneficial a cooling
} system would be. Would a peltier stage be cold enough? Would an LN2
} cold finger be too cold? Thanks.
}
}
} ERiC Jay Miller
} Microscopy & Imaging Specialist
} Electron Probe Instrumentation Center
}
} Northwestern University
} Mail: 2036 Cook Hall
} Office: 1152 Cook Hall
} 2220 Campus Drive
} Evanston, IL 60208-3108
}
} ph: (847) 467-0789
} fax: (847) 467-657
}
} http://www.nuance.northwestern.edu

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

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Email: dridings-at-carolinas.org
Name: Daisy Ridings

Organization: Carolinas Health Care System

Title-Subject: [Filtered] Job oportunity in Charlotte, NC

Message: Carolinas Medical Center in Charlotte, NC is seeking a full-time electron microscopy
technician. The position involves clinical and research work. Duties include operation and basic
maintenance of the transmission electron microscope, sample preparation, thick and thin sectioning,
staining, and digital imaging.

Other required skills include the ability to work independently and interdependently as part of a
high performance team and the ability to troubleshoot problems with techniques and equipment.
Experience with digital imaging and Microsoft Office applications is highly desired.
Immunocytochemistry experience is also helpful.

Bachelors degree required previous TEM experience or electron microscopy certification preferred.

Applicants must apply online at:
http://careers.carolinashealthcare.org/JobDetail.aspx?Pagecode=1&jobid=526543&bID=1702 (direct link).

Carolinas Medical Center is an Academic Medical Center Teaching Hospital and the flagship facility
of Carolinas HealthCare System. Carolinas HealthCare System is an equal-opportunity not-for-profit,
self-supporting public organization offering a wide variety of health and human services to
residents of both North and South Carolina. Carolinas HealthCare System is the largest healthcare
system in the Carolinas, and the third largest public system in the nation.

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Email: cmconway-at-usgs.gov
Name: Carla Conway

Organization: Western Fisheries Research Center

Title-Subject: [Filtered] Zinc formalin as primary fixative for TEM

Message: Hello everyone,

A colleague has tissues which are fixed in zinc formalin and wants to examine them by TEM. Any
thoughts regarding the use of this fixative for EM?

Thanks in advance,

Carla

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} realname - Adrian
} Email - adrian_francisco-at-hotmail.com
} ORGANIZATION - Complutense University of Madrid
} EDUCATION - Graduate College
} LOCATION - SPAIN
} SUBJECT_OF_QUESTION - Corrosion in Metals due to Electron Microscopy
} QUESTION - Dear Microscopist,
}
} Im a student on physics at the Complutense University in Madrid, and
} I have been told by the teacher to write about the negative effects
} on the surface of metals because of the interaction with the
} electrons of the electron beam in microscopy. And how that could be
} a problem at long term range to study the sample.
}
} I'm looking for information on the internet but I dont really go
} anywhere, can you please tell me some specific keywords that I may
} be missing when looking for this??
} Is there any source of information on the internet which talk about
} that particular issue for free?
}
} Thank you very much
}
} Sincerely,
}
} Adrian

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Please start on Google with the search string: "Electron Beam Damage"

Also, in this case, remember that the term "non-destructive" probably originated with a candidate for political office.

Hope this helps,

Fred Monson

Frederick C Monson, PhD
Center for Microanalysis and Imaging, Research and Training (CMIRT)
West Chester University of Pannsylvania
West Chester, PA, 1938
610-738-0437
fmonson-at-wcupa.edu
________________________________________
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From noreply-at-gagnon-inc.com Thu Nov 15 21:15:09 2012
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Dear all

I know that Hitachi H7000 is hardly a new instrument but ours is well maintained and fully operational so we need to consider how best to manage photography in the future.

At present we use Kodak 4489 cut film and it produces excellent results which are then scanned using a flatbed scanner. However there are clearly long term issues about the future of the film, there are the general problems with darkroom processing and chemicals and the inevitable delay in producing results or indeed the cost of staff time if we rush them through.

We are aware of the basic types of digital capture available. In our case 35mm port (wide field capture) or under the film camera (high resolution capture) using something capable of 2k x 2k would be suitable. We mostly image biological tissues and negative stains so the 35mm port would seem sensible.

It would help us greatly if anyone with an H7000 or similar with a retro fitted digital camera system would be willing to share their experiences with us. In particular we would want to know what manufacturer and type,why they chose it, how easy it was to have fitted and how it's worked out. Any info would greatly help us in our decision making.

Thanks

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk
University of Sunderland - Shortlisted for the Times Higher University of the Year 2012

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9, 34 -- Subject: fitting a digital camera to a Hitachi H7000 TEM
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Hi Malcolm

As you may know we run a series of courses known as Protrain Portfolio where
students have teaching data, test specimens and a series of practical
exercises to complete.

These courses go very well when the "student" uses a below screen digital
capture system, but often when using a 35mm port system the resolution is
insufficient to resolve the subtle changes we are trying to have the
students recognise. Remember the 35mm port systems have a much lower
magnification than the below screen systems. But on visiting these
customers we find we are able to resolve the subtle changes on the screen
that were not visible on their 35mm port camera. I do not know the
resolution of the cameras we are taking about, but it worries me that if
people want to see the data they view on the screen the resolution of these
cameras is extremely important, less so for below screen systems.

I should add that some of these customers have been satisfied with their
normal results when I, to be honest, have not!

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: malcolm.haswell-at-sunderland.ac.uk
[mailto:malcolm.haswell-at-sunderland.ac.uk]
Sent: 21 November 2012 14:01
To: protrain-at-emcourses.com

Dear all

I know that Hitachi H7000 is hardly a new instrument but ours is well
maintained and fully operational so we need to consider how best to manage
photography in the future.

At present we use Kodak 4489 cut film and it produces excellent results
which are then scanned using a flatbed scanner. However there are clearly
long term issues about the future of the film, there are the general
problems with darkroom processing and chemicals and the inevitable delay in
producing results or indeed the cost of staff time if we rush them through.

We are aware of the basic types of digital capture available. In our case
35mm port (wide field capture) or under the film camera (high resolution
capture) using something capable of 2k x 2k would be suitable. We mostly
image biological tissues and negative stains so the 35mm port would seem
sensible.

It would help us greatly if anyone with an H7000 or similar with a retro
fitted digital camera system would be willing to share their experiences
with us. In particular we would want to know what manufacturer and type,why
they chose it, how easy it was to have fitted and how it's worked out. Any
info would greatly help us in our decision making.

Thanks

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk
University of Sunderland - Shortlisted for the Times Higher University of
the Year 2012

==============================Original Headers==============================
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9, 34 -- To: "Microscopy MSA (Microscopy-at-microscopy.com)"
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Hi Malcolm:

I too have a Hitachi H-7000 TEM (~25 years old) that have been well maintained. I replaced the film camera with a Gatan MeganScan 4K X 4K digital camera about 10 years ago and I never used the film again.

I have been happy with the Gatan camera, which uses optical fiber (higher sensitivity), instead of glass lenses. It offers high resolution and the software (Digital Micrograph) is easy to use.

The problem with an old TEM is that the camera does not 'talk' to the microscope and you have to input the magnification manually.

The camera is slow in today's standard, and I don't believe Gatan still produces this model anymore.

Hope this helps.

I have no financial interest with Gatan, just a happy user.

Zhaojie

Zhaojie Zhang, Ph. D.
Director, Jenkins Microscopy Facility
University of Wyoming
Laramie, WY 82071
PHONE: 307-766-3038
FAX: 307-766-5625

-----Original Message-----
X-from: malcolm.haswell-at-sunderland.ac.uk [mailto:malcolm.haswell-at-sunderland.ac.uk]
Sent: Wednesday, November 21, 2012 7:06 AM
To: Z.J. Zhang

Dear all

I know that Hitachi H7000 is hardly a new instrument but ours is well maintained and fully operational so we need to consider how best to manage photography in the future.

At present we use Kodak 4489 cut film and it produces excellent results which are then scanned using a flatbed scanner. However there are clearly long term issues about the future of the film, there are the general problems with darkroom processing and chemicals and the inevitable delay in producing results or indeed the cost of staff time if we rush them through.

We are aware of the basic types of digital capture available. In our case 35mm port (wide field capture) or under the film camera (high resolution capture) using something capable of 2k x 2k would be suitable. We mostly image biological tissues and negative stains so the 35mm port would seem sensible.

It would help us greatly if anyone with an H7000 or similar with a retro fitted digital camera system would be willing to share their experiences with us. In particular we would want to know what manufacturer and type,why they chose it, how easy it was to have fitted and how it's worked out. Any info would greatly help us in our decision making.

Thanks

Malcolm

Malcolm Haswell
Imaging Suite
Faculty of Applied Sciences
University of Sunderland
SUNDERLAND
SR1 3SD
UK
email: malcolm.haswell-at-sunderland.ac.uk
University of Sunderland - Shortlisted for the Times Higher University of the Year 2012

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30, 30 -- From ZZhang-at-uwyo.edu Wed Nov 21 09:47:37 2012
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Hi Malcolm,

we own an old Zeiss EM902, which is equipped with a Gatan 694 slow scan
camera (1k x 1k). This one is bottom mounted and we use the viewing
screen to examine the specimen and search for a good position to take a
shot. The search mode of the cam is quite slow (not as slow as our 4k x
4k Ultrascan) and does not replace the viewing screen for this job.
There is no connection to the microscope to read out date. But you can
activate a dialog that asks for the magnification just before you take a
picture.
I think Gatan is quite expensive, but if you can find a used one, the
price can be fine. The Software is only expensive if you need special
plugins for microscope control or EELS.

Michael

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A friend with a Hitachi 5200 wants not only to image long, 3Angstrom wide
lines in glass but to do a compositional analysis. The fact that his
resolution could only ever be an order of magnitude higher even on a metal
sample is not a problem for him, but he would like to get at least to
overcome the charging enough to get a moderately usable resolution. Apart
from use of very low beam energies can anyone offer any tips for this breed
of SEM?

Many thanks for your help!

Ian
Acutance Scientific Ltd

==============================Original Headers==============================
4, 23 -- From ian-at-acutance.co.uk Thu Nov 22 02:50:17 2012
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4, 23 -- From: "Ian Holton" {ian-at-acutance.co.uk}
4, 23 -- To: {Microscopy-at-microscopy.com}
4, 23 -- Subject: Mitigation of charging in glass on Hitachi 5200
4, 23 -- Date: Thu, 22 Nov 2012 08:50:43 -0000
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From noreply-at-musikallegre.com Thu Nov 22 05:05:17 2012
Return-Path: {noreply-at-musikallegre.com}
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Ian,

For imaging and elemental analysis on 3A scale (though I suspect you
meant 3nm) of glass sample I'd suggest your friend to go through the
pain of preparing a lamela and trying FE TEM with EELS at friendly
University facility. There is some hope that electron beam induced
conductivity will help mitigate charging in the glass...

Or maybe try gentle coating by Iridium in turbo-pumped sputter coater?

But elemental analysis on 3A, or even 3nm scale, in even the best SEM
may turn out to be too much...

Cheers :)

Valery Ray
=================================
PBS&T, MEO Engineering Co., Inc.
290 Broadway, Suite 298
Methuen, MA 01844 USA
Phone: +1-978-296-5063 - leave a message with call-back number
US Mobile: +1-978-305-0479
Skype: pbstmeo
E-mail: vray-at-partbeamsystech.com
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Web: www.freudlabs.com

On 11/22/2012 3:51 AM, ian-at-acutance.co.uk wrote:
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} A friend with a Hitachi 5200 wants not only to image long, 3Angstrom wide
} lines in glass but to do a compositional analysis. The fact that his
} resolution could only ever be an order of magnitude higher even on a metal
} sample is not a problem for him, but he would like to get at least to
} overcome the charging enough to get a moderately usable resolution. Apart
} from use of very low beam energies can anyone offer any tips for this breed
} of SEM?
}
} Many thanks for your help!
}
} Ian
} Acutance Scientific Ltd
}
}
} ==============================Original Headers==============================
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} 4, 23 -- To: {Microscopy-at-microscopy.com}
} 4, 23 -- Subject: Mitigation of charging in glass on Hitachi 5200
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Title-Subject: [Filtered] 46th Annual NESM Fall Symposium & Business Meeting - Nov. 30th

Message: Join NESM for our 46th Annual Fall Symposium and Business Meeting hosted by Gordon College
on Nov. 30, 2012. The meeting is composed of five technical talks, a business meeting, and a buffet
dinner. Please check out our website for more information:
http://nesmicroscopy.org/upcoming-meeting.html

12:30 - Meeting Registration
1:00 - Welcoming Remarks
1:10 - Neurogenesis in adult brains: A hematopoietic connection?, Barbara S. Beltz, Ph.D.,
Jeanne L. Benton and Paula G. Chaves da Silva, Wellesley College, Wellesley, MA
2:00 - Printing at the micro and nano scale, Alexander Smetana, Ph.D., NanoInk Inc.
2:50 - Coffee Break
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Email: Pinhead01-at-msn.com
Name: Pierre Bustanoby

Organization: Specialized Electron Microscope Service

Title-Subject: [Filtered] SEM Imaging

Message: Good morning!

I'm looking for any 4PI computer interface board.

Either PC or Mac would be wonderful.

Actually......... I'm interested in any image capture system that would work
On a JEOL 840.

Please contact:

Pierre Bustanoby
SEMS
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In the Pacific Northwest
206 795-1599

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Name: Ravi Thakkar

Organization: NIMHANS, Bangalore, INDIA

Title-Subject: [Filtered] Elemental Analysis of Metabolic inclusions.

Message: Dear Listener,
Does any one has experience on elemental analysis of Metabolic inclusions (Metabolic storage
disorder) and defined its molecular architecture.

Kindly share your experience.

Thanks.

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Hi

3Angstrom from an SEM, I doubt if you will get very far, perhaps you mean
3nm? To carry out elemental analysis at this level to obtain any
discrimination you need STEM, OK that includes STEM in a SEM.

We have worked on a FEGSEM (30kV) mapping at 500,000X but only by using a
ultramicrotome to prepare the specimen; we were mapping 5nm particles on a
reasonably friendly specimen. Even in a Lab6 SEM (40kV) using the same
technique we were able to map particles below 0.1micron

I would be very interested on other peoples comments?

Steve

Steve Chapman FRMS
Protrain for Consultancy and Courses in Electron Microscopy
Tel & Fax +44 (0)1280 816512 Cell +44 (0)7711 606967
www.emcourses.com

-----Original Message-----
X-from: ian-at-acutance.co.uk [mailto:ian-at-acutance.co.uk]
Sent: 22 November 2012 08:51
To: protrain-at-emcourses.com

A friend with a Hitachi 5200 wants not only to image long, 3Angstrom wide
lines in glass but to do a compositional analysis. The fact that his
resolution could only ever be an order of magnitude higher even on a metal
sample is not a problem for him, but he would like to get at least to
overcome the charging enough to get a moderately usable resolution. Apart
from use of very low beam energies can anyone offer any tips for this breed
of SEM?

Many thanks for your help!

Ian
Acutance Scientific Ltd

==============================Original Headers==============================
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glass on Hitachi 5200 4, 23 -- Date: Thu, 22 Nov 2012 08:50:43 -0000 4, 23
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Thanks for the replies. I expressed myself very badly re the resolution. My
intended meaning was that it does not matter for him to get resolution
anywhere close to the 3Angstroms line-width or even 3nm or 30nm as long as
he can get some compositional analysis. For instance if the 3A wide lines
appeared (due to limitations of instrumental resolution and charging) to be
factors worse, this would not matter in itself if he could get one
measurement of the composition, maybe as a small peak on a huge background.

However, I see the clincher has to be that in any case, the ratio of
interaction volume to material in the lines is too large to get a meaningful
measurement.

Many thanks for your comments!

Ian

Ian Holton
Acutance Scientific

-----Original Message-----
X-from: ian-at-acutance.co.uk [mailto:ian-at-acutance.co.uk]
Sent: 22 November 2012 09:01
To: ian-at-acutance.co.uk

A friend with a Hitachi 5200 wants not only to image long, 3Angstrom wide
lines in glass but to do a compositional analysis. The fact that his
resolution could only ever be an order of magnitude higher even on a metal
sample is not a problem for him, but he would like to get at least to
overcome the charging enough to get a moderately usable resolution. Apart
from use of very low beam energies can anyone offer any tips for this breed
of SEM?

Many thanks for your help!

Ian
Acutance Scientific Ltd

==============================Original Headers==============================
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Hi all,

This may or may not be a trivial question. I have been given some
samples of particles suspended in water with a little SDS/SLS (sodium
dodecyl sulphate) surfactant thoughtfully added to keep them from
settling. The general plan is to catch some particles on a carbon grid
as normal, but I would like a little advice to avoid disaster, since I
only have a few ml of suspension to try things out with.

So - has anybody made a powder sample from a solution with SDS in?

Does it result in a horrible contaminating organic mess on the sample
or is it a perfectly reasonable thing to pipette straight from such a
liquid onto the grids just like you would for e.g. isopropanol?

If it would be best to remove the SDS before making powder samples,
does anybody have a better method than pouring the suspension through
a filter paper and washing the solid off the paper with a more
sensible liquid?

Hoping I don't look too silly

Jo Sharp, Sheffield

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7, 37 -- Subject: TEM samples from SDS solution
7, 37 -- From: Joanne Sharp {j.sharp-at-sheffield.ac.uk}
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Hi Jo,

Are the particles charged? You might want to look into coating grids with a water soluble poly-amino acid (or other simple polymer) with opposite charge. E.g. gold particles (-) easily bind to grids coated with poly-L-lysine (+) in a wide pH range.

Sulphate would certainly interfere somewhat, but it might be worth a try.

The procedure is very simple and gives neat results with the least possible contamination. If you need more info, contact me off list, please.

Good luck,

Jan Leunissen
Aurion

i: www.aurion.nl

On 24/11/2012, at 5:08 AM, j.sharp-at-sheffield.ac.uk wrote:

}
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} Hi all,
}
} This may or may not be a trivial question. I have been given some
} samples of particles suspended in water with a little SDS/SLS (sodium
} dodecyl sulphate) surfactant thoughtfully added to keep them from
} settling. The general plan is to catch some particles on a carbon grid
} as normal, but I would like a little advice to avoid disaster, since I
} only have a few ml of suspension to try things out with.
}
} So - has anybody made a powder sample from a solution with SDS in?
}
} Does it result in a horrible contaminating organic mess on the sample
} or is it a perfectly reasonable thing to pipette straight from such a
} liquid onto the grids just like you would for e.g. isopropanol?
}
} If it would be best to remove the SDS before making powder samples,
} does anybody have a better method than pouring the suspension through
} a filter paper and washing the solid off the paper with a more
} sensible liquid?
}
} Hoping I don't look too silly
}
} Jo Sharp, Sheffield
}
} ==============================Original Headers==============================
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} 7, 37 -- Subject: TEM samples from SDS solution
} 7, 37 -- From: Joanne Sharp {j.sharp-at-sheffield.ac.uk}
} 7, 37 -- To: Microscopy-at-microscopy.com
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--Please respect nature in and around New Zealand's precious waterways and refrain from jet boating--

==============================Original Headers==============================
18, 21 -- From leunissen-at-aurion.nl Fri Nov 23 20:03:41 2012
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Howdy Y'all,

This is not a confocal related question. BUT since this forum is such a
big group of microscopists, I'd figure I would try anyway...

I am currently trying to find a PDF version of the Hamamatsu Orca ER user
guide/manual. Does anyone have access to it?! Searches on Google only
brought up the datasheet.

Thanks for any and all help in advance!!

Regards,
Pete

--
Peter Gabriel Pitrone - TechRMS
Microscopy/Imaging Specialist
Prof. Dr. Pavel Tomancak group
Max Planck Institute for
Molecular Biology and Genetics
Pfotenhauerstr. 108
01307 Dresden

"If a straight line fit is required, obtain only two data points." - Anon.

==============================Original Headers==============================
10, 29 -- From pitrone-at-mpi-cbg.de Mon Nov 26 05:34:02 2012
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Hi Jo!
�
"only" a few ml for an electron microscopist is a lot!!
You probably only need 5-10�l per grid so it's not a big deal to test it.
This easiest thing I can think of to get rid off SDS is simply dialysis against water.
The biggest deal will be to translate kDa (usually used to calibrate the dialysis bag) in nm/�m but I am sure that google has the answer.
Have fun.
�
Stephane
�

X-from: "j.sharp-at-sheffield.ac.uk" {j.sharp-at-sheffield.ac.uk}
To: nizets2-at-yahoo.com
Sent: Friday, November 23, 2012 5:12 PM

Hi all,

This may or may not be a trivial question. I have been given some
samples of particles suspended in water with a little SDS/SLS (sodium
dodecyl sulphate) surfactant thoughtfully added to keep them from
settling. The general plan is to catch some particles on a carbon grid
as normal, but I would like a little advice to avoid disaster, since I
only have a few ml of suspension to try things out with.

So - has anybody made a powder sample from a solution with SDS in?

Does it result in a horrible contaminating organic mess on the sample
or is it a perfectly reasonable thing to pipette straight from such a
liquid onto the grids just like you would for e.g. isopropanol?

If it would be best to remove the SDS before making powder samples,
does anybody have a better method than pouring the suspension through
a filter paper and washing the solid off the paper with a more
sensible liquid?

Hoping I don't look too silly

Jo Sharp, Sheffield

==============================Original Headers==============================
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Dear All,

Does anyone have experience funding short research visits (3 months)
to Germany (specifically, the Max Plank Institute) from within the EU?

Many thanks for any help you can provide me.

Regards,

Arturo Tejada

--

===================================================
Dr. Arturo Tejada
Assistant Professor
Mechatronic System Design Group
Precision and Microsystems Engineering Dpt.
Mekelweg 2, Room G-1-300
2628 CD, Delft, The Netherlands
Phone: +31-15-27-81550
Fax: +31-15-27 89475
www.tejadaruiz.net
===================================================

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Malcolm
I have a Hitachi 7100 TEM (basically a 7000 with a motorized stage)
and had a Gatan 7 megapixel bottom mount camera installed about 5 years
ago. The installation went smoothly and we have been very happy with the
high resolution images which have been used in a number of
publications. The digital micrograph software is easy to use and no one
in our lab has any desire to go back to film. We do mostly bacteria and
viral imaging so the bottom mount camera made sense for us.

Tony Greco

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If you're a non-profit, you're still using film, and you're willing to
pay for shipping.
They're as old as 1988 and as young as 10/89, and have been refrigerated
from purchase (except for their brief time being shipped to me about 8
years ago, but then, there were twice as many boxes). The film was
still fine in January of this year.
We've finally gotten all of the users onto the new digital camera on our
old EM-10C.
Julian

--
Julian P.S. Smith III
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Dept. of Biology
Winthrop University
349 Columbia Ave
Rock Hill, SC 29733

803-323-2111 x6427 (vox)
803-323-3448 (fax)
803-524-2347 (cell)
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Personal Website www.rociada-east.net

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4, 16 -- Subject: 12 boxes of 4489 EM film, 3 1/4" x 4"
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Thanks, all.
Julian

--
Julian P.S. Smith III
Director, Winthrop Microscopy Facility
Dept. of Biology
Winthrop University
349 Columbia Ave
Rock Hill, SC 29733

803-323-2111 x6427 (vox)
803-323-3448 (fax)
803-524-2347 (cell)
Research Website www.birdnest.org/smithj
Personal Website www.rociada-east.net

==============================Original Headers==============================
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Email: innap-at-savion.huji.ac.il
Name: Inna Popov

Organization: The Hebrew University of Jerusalem

Title-Subject: [Filtered] TEM samples from SDS solution

Message: Hi,
A reasonably good TEM sample could be prepared from 2-3 microliter of suspension. If your NPs are
heavy and dense enough (i.e. an inorganic staff)and their content is not extremetly low you will be
able to identify them on an amoprhous carbon support even with SDS presenting in the sample.
} From my experience, under primarily drying ( you may do that in any vacuum chamber pumped with rotary pump) a liquid structure is broken and the surfactant is spontaneously separated from other components of dispersion. At high content it may solidify aside as semi-transparent body on top of your grid so that you may remove it from grid by twizers. At lower content SDS may crystallize in micron-scale surface crystallites on a grid apart from NPs. At very low content you will observe 3D amorphous low contrast features on a grid still apart from your NPs. So, you see that SDS will most likely not disturb your imaging except of one sad case of expected organic nanoparticles in dispersion. In this case I would never dry the dispersion at all, but go to any cryo-EM technique preserving the structure of liquid. Otherwise you would most likely loose the true morphology and size of your organic NPs.
Good luck,
Inna

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Email: vakimler-at-med.wayne.edu
Name: Vickie Kimler

Organization: Wayne State University - School of Medicine

Title-Subject: [Filtered] focused ion beam - biological applications

Message: Hello,
Does anyone out there have any insights on focused ion beam for biological specimens in Durcupan
embedment medium? We are doing a pilot at a neighboring university and are trying to image mouse
cardiac tissue. The tissue was en bloc stained with 2% OsO4 in 0.12 M cacodylate buffer, followed by
30% ethanolic 2% uranyl acetate and then dehydrated. I am preparing a block face by diamond knife
ultramicrotomy, but I need to know the size of the face.

The tissue chunks are about 2-3 mm in either dimension.
TEM ascertained they are well preserved.

I will cut off the block base so the conical tip with the apical block face remains, about 5 mm in
height, and no greater than that. Then this tip will be mounted on a metallic agent covered stub to
reduce charging.

Do I then coat the entire sample in gold, at about 25 nm?

Thanks; any other tips would be helpful.
Vickie

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Dear Vickie,

Here is a link to a publication that we tried to follow
(http://www.frontiersin.org/Neuroanatomy/10.3389/neuro.05.018.2009/full).
Our collaboration efforts ended without success, but more due to external
factors (lack of FIB experience in bio samples, great distance from machine,
etc) that to errors in methodology. Hope you can have more success when you
have your machine nearby.
As much as I understood, in Zeiss' FIB you can mount a much bigger sample
than those 5x5mm.

Wish you luck!
Josif

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

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Email: vakimler-at-med.wayne.edu
Name: Vickie Kimler

Organization: Wayne State University - School of Medicine

Title-Subject: [Filtered] focused ion beam - biological applications

Message: Hello,
Does anyone out there have any insights on focused ion beam for biological
specimens in Durcupan embedment medium? We are doing a pilot at a
neighboring university and are trying to image mouse cardiac tissue. The
tissue was en bloc stained with 2% OsO4 in 0.12 M cacodylate buffer,
followed by 30% ethanolic 2% uranyl acetate and then dehydrated. I am
preparing a block face by diamond knife ultramicrotomy, but I need to know
the size of the face.

The tissue chunks are about 2-3 mm in either dimension.
TEM ascertained they are well preserved.

I will cut off the block base so the conical tip with the apical block face
remains, about 5 mm in height, and no greater than that. Then this tip will
be mounted on a metallic agent covered stub to reduce charging.

Do I then coat the entire sample in gold, at about 25 nm?

Thanks; any other tips would be helpful.
Vickie

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Email: vladimir.girman-at-upjs.sk
Name: Vladimir Girman

Organization: Department of Solid State Physics - University of P.J.Safarik in Kosice

Title-Subject: [Filtered] Gun alignment

Message: Hi everyone,

do have someone smart guideline hints to gun alignment on JEOL 2100F. If I use a gun wobblers
(shift, tilt or angle) I see only streak and I don�t know what to do with it. When it is well
aligned? What I have to do with wobbled gun spot? And how to recognize the gun is not in optical axis?

Many thanks for your advices
Vladimir Girman

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Email: ravi.thakkar369-at-gmail.com
Name: Ravi Thakkar

Title-Subject: [Filtered] Elemental Analysis of Metabolic inclusions in lysosomal storage disorder.

Message: Dear Listener,
My previous post is not much precise, so I would like to elaborate my query.
Does any one have experience on Elemental analysis of metabolic inclusions (Lysosomal Storage
Disorder)in human tissue using Electron microscopy and defined chemical composition of complex
metabolites form inclusions and its morphological study.

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Email: john.mitchels-at-gmail.com
Name: john Mitchels

Organization: University of Bath, UK

Title-Subject: [Filtered] 1 Day Microscopy Conference University of Bath

Dear Listers

The University of Bath in the UK will be holding a 1 day mini conference in
Microscopy and Spectroscopy on the 9th Januray 2013.

Please see this weblink for the pdf programme
https://www.dropbox.com/s/nqpddtlhoew44a5MAS%20Conference%20Program%202013=.pdf

The focus of this meeting is to bring together
people from all disciplines from the south west interested in microscopy and spectroscopy. It is a
great chance to meet people who work in similar fields in the area and to see some of the exciting
latest developements.

This year we have another exciting lineup of speakers with wide ranging techniques including TEM,
SEM, AFM, XRD, Raman, Confocal, CT tomography which span the disciplines.

It is open to all. University based reseachers can attend free and our corporate brothers and
sisters can attend for 30 ukp per person. There is a student poster session with prizes.

There will be trade stands from selected companies.

If you are intersested in attending please email mas-at-bath.ac.uk quoting
'conference 2013'.

I look forward to meeting you here.

Dr John Mitchels
Microscopy and Analysis Suite
University of Bath, UK

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Hi

I am about to start using Quetol 651 resin on an ultramicrotome..
Have never used this before and the information with the kit is limited.
I would appreciate any tips anyone can provide.

What is the best way to collect the sections?
Dry?
Float on water or some other liquid?

Slides:
Collect onto plain slides or electrostatic ones?
Perhaps some other coating?

Dry them at room temperature over night?
Dry them in a 60 deg C oven?

Regards

Leigh Silvester
This message and any attachment are intended solely for the addressee and may contain confidential information. If you have received this message in error, please send it back to me, and immediately delete it. Please do not use, copy or disclose the information contained in this message or in any attachment. Any views or opinions expressed by the author of this email do not necessarily reflect the views of the University of Nottingham.

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Email: amit.welcomes.u-at-gmail.com
Name: Amit Gupta

Organization: IISc

Title-Subject: [Filtered] Determination of Cs

Message: Good morning everyone,
We are using JEOL 2100f. We are trying to characterize the whole instruement. For that i need to
find out spherical aberration constant. I used the method used in JC Spence. but problem is that
when i give the beam tilt, the image gets too astigmatic, am i doing something wrong?
i tried correcting astigmatism and took images again, but the Cs came out to be 7 x 10^-9!!!
Can anyone shed any light on this?

Thank You

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Registration is open for The Histochemical Society Immunohistochemistry
and Microscopy Course, now in its fourth year at the Marine Biological
Laboratory in Woods Hole, MA from March 9 - 14, 2013 and we include a
brief description of the course below. However, we also want to update
you on Travel Awards available for the course through HCS and the FASEB
MARC Program.

HCS is offering Student Travel Awards to offset the cost of attending
the HCS IHCM Course. The awards are based on how the course will benefit
you in your work. The application deadline is January 18, 2013.
Information about the HCS Travel Awards can be found on the the HCS website:
http://histochemicalsociety.org/Awards/Students---Post-Docs.aspx

Additionally, students from groups underrepresented in science may apply
for financial support for travel-related expenses and course
registration through the FASEB MARC Travel Program. The DEADLINE for
submission is February 12, 2013. These awards are processed on a first
come basis. Information about the program may be found on their website:
http://www.faseb.org/MARC-and-Professional-Development/Travel-Awards/Scientific-Meetings-and-Conferences.aspx

You may access the MARC application here:
(http://bit.ly/IHCM2013TravelAward).

VERY IMPORTANT: You have until the deadlines listed above to apply for
awards but YOU MUST REGISTER FOR THE COURSE BY JANUARY 10, 2013

ABOUT THE COURSE:
The Immunohistochemistry and Microscopy (IHCM) course provides
participants in-depth theory of and extensive hands-on experience with
immunohistochemistry (IHC) techniques as well as theory and hands-on
experience with a broad range of microscopic imaging techniques. The
course emphasizes hands-on experience, troubleshooting, and the exchange
of ideas between and among faculty and participants.

The course will enable participants to independently carry out IHC and
imaging in their laboratories and to critically evaluate and
troubleshoot problems using IHC and microscopy techniques. The
interplay of theory, hands-on experience, and discussion is central to
this course that serves advanced undergraduates, graduate students,
laboratory technicians, postdoctoral students, and new and established
faculty/research clinicians.

The course is limited to 30 students and registration includes
accommodations and all meals and breaks. Accommodations are on the
campus of MBL. We look forward to seeing you in March.

Eduardo Rosa-Molinar, Ph.D.
Group Leader, Biological Imaging Group (BIG)
Associate Professor of Biology and Neurobiology
University of Puerto Rico-Rio Piedras
P.O. Box 21809 UPR Station
San Juan, Puerto Rico 00931-1809
Phone: (787) 764-0000; Ext. 1-2850
Facsimille: (888) 762-7577
Webpage: http://pisces.cnnet.clu.edu/erm-lab/

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Email: amit.welcomes.u-at-gmail.com
Name: Amit Gupta

Organization: IISc

Title-Subject: [Filtered] Focal series reconstruction

Message: Good morning,
We have a Jeol 2100f microscope with olympus keenview ccd camera. We want to perform EWR. But
unfortunately neither jeol seems to have any package like TrueImage by fei, neither olympus any
plug in like that for Gatan Digital EM.
Does any one have any alternative?
Has anyone tried it themselves using Matlab etc? (it has been done before as publications exist)

Thank You

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Email: dan.buntman-at-wdc.com
Name: Dan Buntman

Organization: Western Digital

Title-Subject: [Filtered] Sr. TEM engineer position

Message: We currently have an open position for a Sr. Principal Engineer-TEM Specialist in the
Analytical Services Group at Western Digital Media R & D. The job postings for all N. American
positions canbe found at: http://www.wdc.com/en/company/employment/northamerica.aspx.

Alternatively you can submit your resume to dan.buntman-at-wdc.com.

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Email: xulingling723-at-gmail.com
Name: Lingling

Organization: Dalhousie university

Title-Subject: [Filtered] Defect of video of a motion

Message: Hello,

When I'm using ZEISS AXIOVERT 200M INVERTED MICROSCOPE to record a process of fiber being pulled
from a viscous solution, there's always a defect of a black/white spot (either really black or very
bright depending on the focus)at the spot where the viscous solution was lifted up by the fiber that
formed "convex" structure. Does anyone know what caused the black/white spot?

Thanks in advance,

Lingling

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Dear Colleagues,

It is a pleasure to invite you to participate in the 16^th International
Conference on Noncontact Atomic Force Microscopy (NC-AFM 2013) in
conjunction with the symposium on �Atomic Force Microscopy for Energy
Applications". These events will be organized by the University of
Maryland, College Park, and the National Institute of Standards and
Technology (NIST), and will take place at the University of Maryland on
August 5-9 of 2013.

The NC-AFM 2013 meeting follows a successful series of international
conferences devoted to the latest progress in dynamic atomic force
microscopy (AFM). The focus is on experimental, theoretical, and
instrumental developments in frequency modulation and other dynamic
operation modes with particular emphasis on high-resolution imaging and
force spectroscopy. As has become tradition, there will be a
PRE-CONFERENCE SYMPOSIUM which will have the title �Atomic Force
Microscopy for Energy Applications� (described below).

We welcome contributions for oral and poster presentations on the
following topics:

�Novel instrumentation and techniques in AFM

�Atomic resolution imaging on insulating substrates, semiconductors, and
metals

�High-resolution imaging of molecules, clusters and biological systems

�Atomic and molecular-scale manipulation

�Simultaneous force and tunneling spectroscopy

�High-resolution imaging and spectroscopy in liquid environments

�Theoretical analysis of contrast mechanisms, forces and tunneling
phenomena

�2D and 3D force-field mapping techniques

�Small amplitude and lateral force measurements using dynamic methods

�Mechanisms and understanding of damping and energy dissipation

�Nanoscale measurements of charges, work function, and magnetic properties

�Theoretical aspects of scanning probe techniques

�Tapping mode versus non-contact mode imaging

�Multi-frequency and spectral AFM

SYMPOSIUM (CONFERENCE DAY 1): The symposium �Atomic Force Microscopy for
Energy Applications,� chaired by Sergei Kalinin (Oak Ridge National
Laboratory, USA), will cover recent advances in characterization of
energy relevant materials using advanced AFM methods, including
applications in which AFM is combined with other techniques for
measurement and modification. Relevant applications include KPFM, ESM
and CAFM of photovoltaic materials, electrochemical imaging and
tip-induced electrochemical reactions, electric imaging in liquids and
imaging of electrochemical processes. Submissions are also sought which
highlight novel dynamic AFM modes and platforms tailored specifically
for energy materials, measurements under in-situ and in-operando
conditions, combined AFM-optical, STEM, and X-ray studies, as well as
novel approaches for SPM data analysis and image quantification for
extracting energy-relevant functionalities. Emphasis on the elucidation
of energy transformation pathways and fundamental physical and chemical
mechanisms involved in energy conversion and storage in all types of
environments is also desired.

PROCEEDINGS: The proceedings will be published in the Beilstein Journal
of Nanotechnology (open access with no publication charges:
https://www.beilstein-journals.org/bjnano/home/home.htm) and will be
edited by Udo Schwarz (Yale University, USA) and Mehmet Baykara (Bilkent
University, Turkey).

CONFERENCE WEBSITE: A webpage in construction is available at
http://meetings.umd.edu/ncafm2013

Further information and instructions will be posted on this website as
they become available.

DEADLINES:

Abstracts due April 22

Acceptance notification May 20

Early registration ends June 24

Proceedings deadline October 14

We look forward to your submissions.

Sincerely,

The organizing Committee:

Santiago Solares, University of Maryland (UMD) � Conference Chairman

William Cullen, UMD

Michael Dreyer, UMD

Joseph Stroscio, NIST

Robert Cook, NIST

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Hello,

Sorry for the delay. And thanks to all that shared they experience with me. It was really useful. I was busy and was trying to put things together. I got a few tips about dual-beam from FEI and Zeiss. And I also did talk to some of the representatives from FEI, Zeiss and Tescan in Brazil. Why these three? They have a local service team. All agree that none of the systems on the market are perfect and have their problems. Someone mentioned that a Zeiss dual-beam took 9 months to be installed. But once it was working, it was a great instrument. And you get a Gemini column with it. Of coarse, not exactly the same from SEMs for design reasons. About FEI, someone complained about how expensive their service contract is and how they increased it every year. We had two FEG-SEMs from FEI installed in our lab a year ago. We regret about choosing them. Both instruments (Inspect and Quanta) are plagued with some design problems. Detectors (BS) that break apart on the pole piece. Touch detector that do not detect when the stage touches detectors (EBSD). To mention a few. I can't say much about Zeiss since we don't have one. But someone in Brazil was complaining about one specific service engineer from Zeiss creating new problems instead of fixing the original problem.

Some people suggested to try Tescan. I talked with them and I find out that they are located near to the FEI facility in Brno, Czech Republic. I was told that they share/exchange engineers. The thing interesting is that Tescan and Zeiss uses the same FIB column from Orsay Physics. The one thing we liked was that Tescan offered to us a ToF-SIMS for the dual-beam. Zeiss have something similar, but the engineer we talked to didn't know much since it is new. If someone is interested, Tescan send me a paper about it:

"High spatial resolution time-of-filght secondary ion mass spectrometer for the masses: a novel orthogonal ToF FIB-SIMS instrument with in situ AFM", J.A. Whitby et al., Adv. in Mat. Sci. and Technol. Vol. 2012, Article ID 180437.

We are still adjusting the package for our dual-beam. Right now, Zeiss is offering the best deal. That was very surprising to us. Last year, at the time we went for new SEMs, Zeiss was not much motivated. We are still negotiating and planning to do the sample prep with all dual-beam we can get our hands on. Probably after finalizing the configuration we want, in February time frame. Then we can learn if there are differences in the lift-out process for each machine.

Carlos

__
Carlos Kazuo Inoki
Brazilian Nanotechnology National Laboratory
Caixa Postal 6192
Campinas, SP
13083-970
Brazil

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Sent: Saturday, November 10, 2012 00:56
To: Roy Golombick

Hello,

We are evaluating to purchase a dual-beam FIB for our lab. It is going
to be used mainly for TEM sample preparation (semiconductor, ceramics,
metal, soft...). Looking for the options available in the market, we
narrow it down to FEI and Zeiss. Probably FEI Nanolab or Zeiss Auriga.
The kind of information I am looking is about what is your experience
with these instruments. Specially maintenance and how much if costs per
year in real world (parts and service).

One advantage about Zeiss was the Argon beam for sample cleaning. But I
was told that it is not available for the Auriga dual-beam and It is not
really necessary to prepare good TEM samples. Tripod polishing produces
wonderful samples, but most people have some difficulty in doing so. Did
you stop using tripod polishing after start using FIB? Does low energy
ion milling helps to improve the quality of FIBed samples (tripod
quality)?

Any opinion or experience that you want to share will be very useful
for us.

Thanks,

Carlos
____________________________________________
Carlos Kazuo Inoki
Brazilian Nanotechnology National Laboratory Rua Giuseppe Maximo
Scolfaro 10000 Campinas, SP 13083-970 Brazil

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Having worked on mucosal tissues most of my professional career, I think I know the answer to my own question but I can't prove it! �

I have always used the phrase "lamina propria" to refer to the connective tissue layer below the epithelium and above the muscularis mucosae. I have assumed the phrase "substantia propria" is for a similar layer but more correctly used for the connective tissue layer below an epithelium that doesn't have a muscularis mucosae and therefore no distinction between a lamina propria and underlying submucosa. But despite aggressive Googling, I am unable to find a definition of substantia propria from an authoritative source. Is this something I made up in my head (perhaps they are true synonyms that reflect country of use) or is my definition precise? Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Email: williams-at-genectr.hunter.cuny.edu
Name: Lloyd Williams

Organization: Hunter College

Title-Subject: [Filtered] Opening To Teach TEM Course at Hunter College

Message: Dear List
Hunter College is looking for an individual to teach a one semester electron microscopy course in
the Spring of 2013. The school is located on the upper east side of New York City, at 68th street
and Lexington. The class would meet twice a week for lectures and lab. The class meeting are
currently scheduled for Tuesday and Wednesday afternoon.
It would be at the introductory level covering the basic
theoretical background of TEM, sample preparation and imaging. The position would be at the level of
an adjunct faculty appointment. Interested candidates should email their resume to
microscopy-at-genectr.hunter.cuny.edu

Thank you

Dr. Lloyd Williams
Dept. of Biology
Hunter College
695 Park Ave
New York NY 10065

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http://education.yahoo.com/reference/gray/subjects/subject/225 [cornea]

http://education.yahoo.com/reference/gray/illustrations/figure?id=871

http://medical-dictionary.thefreedictionary.com/Substantia+propria [synonyms at bottom of paragraph]

I remember the rumor, years ago, that Harvard Med, for a brief moment, declared 'gross' anatomy for medical students to be optional. Hmmmm!

I hear complaints from academia on high that there are no histologists to analyze research specimens. If there are none, then there can be no trainers, expecially if the 'specialty' has been labelled archaic. As I peruse the current hardbacks in histology, they are found to be directed to pathologists. Histology has become relegated to soft cover bindings.

Nomenclature is a pain. Thus, there are those who spend their careers relegating old terminology to oblivion by not learning it in the first place. In this case, it turns out that 'substantia propria' was used to describe poorly fixed, compact material in the cornea that lay between layers #2 and #4 of 5 of Grey's Anatomy, Fig. 871 [cited above].

It is not lamina propria, and it would be appropriate to print this response, file it, and spend a few evenings at the local pub trying to target that part of the neural net that holds the term 'substantia'. Thus, will 'propria' be retained unencumbered with an archaic Latin term whose meaning in Emglish is, at best, vague - at least according to 1 or 2 of the last Latin experts in the world.

I hope all of this helps, because you and I, Prof Phillips, will likely be the last of our kind to discuss this part of histologic anatomy.

Cheers, and Merry Christmas to all,

Fred Monson

P.S. Maximow, Bloom, Ham and Fawcett. Those from whose hard-bound books I learned, and then, taught this stuff.

Frederick C Monson, PhD
Center for Microanalysis and Imaging, Research and Training (CMIRT)
West Chester University of Pannsylvania
West Chester, PA, 1938
610-738-0437
fmonson-at-wcupa.edu
________________________________________
X-from: PhillipsT-at-missouri.edu [PhillipsT-at-missouri.edu]
Sent: Thursday, December 06, 2012 6:55 PM
To: Monson, Frederick

Having worked on mucosal tissues most of my professional career, I think I know the answer to my own question but I can't prove it!

I have always used the phrase "lamina propria" to refer to the connective tissue layer below the epithelium and above the muscularis mucosae. I have assumed the phrase "substantia propria" is for a similar layer but more correctly used for the connective tissue layer below an epithelium that doesn't have a muscularis mucosae and therefore no distinction between a lamina propria and underlying submucosa. But despite aggressive Googling, I am unable to find a definition of substantia propria from an authoritative source. Is this something I made up in my head (perhaps they are true synonyms that reflect country of use) or is my definition precise? Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Dear Listers,
I'm quite embarassed asking your help just for setting up Koehler
condition in my Leica DM LM. I studied it a long time ago but I've never
needed to use it. Now it's time, but I cannot manage to get the right
condition.

With the specimen in focus, field aperture rim appears quite blurred
becuase of chromatic aberration, but maybe it's normal given the
condenser is just an achromat (CL/PH 0.90/1.25 S1); and it looks almost
in focus only if condenser aperture is almost closed and the condenser
is at the end of its focus range, by the sample side.
Besides I'm not able to find the filament image in objective bfp,
watching in the tube without eyepieces while moving the condenser along
its whole focus range. Maybe this image is just so demagnified that I
didn't realize its presence.

Could any one give me some very basic hint?
Thanks a lot.

Davide Cristofori

~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~
Davide Cristofori

direct phone = 041.234.67.26
e-mail = dcristofori[at]unive[dot]it

Universit� Ca' Foscari Venezia
Dipartimento di Sc. Molecolari e Nanosistemi
Via Torino, 155b I-30172 Mestre (VE) Italy
~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~

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I appreciate the thoughts but not remained less than fully satisfied. The substantia propria is also used to describe connective tissue in the conjunctiva and sclera, in the ear in relation to the membrane tympani and less commonly in the uterus. The citation from Gray's on cornea might indeed be an early "defining moment" that entrenched the use of this term for mostly ocular tissues. My angst over using it results mostly from not understanding why a novel term is used here. Was there a unique feature (lamellated fibers?) that makes it a substantia propria or simply a historical practice for those in the ocular milieu. I can't speak to Harvard's practice on teaching gross anatomy but when I played a small part in teaching Histology there, the course was legendary for the quality of job done by Marian Neutra and Dan Goodenough. Sadly, as in many Histology classes, ocular tissues weren't discussed.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: FMonson-at-wcupa.edu [mailto:FMonson-at-wcupa.edu]
Sent: Thursday, December 06, 2012 7:39 PM
To: Phillips, Thomas E.

http://education.yahoo.com/reference/gray/subjects/subject/225 [cornea]

http://education.yahoo.com/reference/gray/illustrations/figure?id=871

http://medical-dictionary.thefreedictionary.com/Substantia+propria [synonyms at bottom of paragraph]

I remember the rumor, years ago, that Harvard Med, for a brief moment, declared 'gross' anatomy for medical students to be optional. Hmmmm!

I hear complaints from academia on high that there are no histologists to analyze research specimens. If there are none, then there can be no trainers, expecially if the 'specialty' has been labelled archaic. As I peruse the current hardbacks in histology, they are found to be directed to pathologists. Histology has become relegated to soft cover bindings.

Nomenclature is a pain. Thus, there are those who spend their careers relegating old terminology to oblivion by not learning it in the first place. In this case, it turns out that 'substantia propria' was used to describe poorly fixed, compact material in the cornea that lay between layers #2 and #4 of 5 of Grey's Anatomy, Fig. 871 [cited above].

It is not lamina propria, and it would be appropriate to print this response, file it, and spend a few evenings at the local pub trying to target that part of the neural net that holds the term 'substantia'. Thus, will 'propria' be retained unencumbered with an archaic Latin term whose meaning in Emglish is, at best, vague - at least according to 1 or 2 of the last Latin experts in the world.

I hope all of this helps, because you and I, Prof Phillips, will likely be the last of our kind to discuss this part of histologic anatomy.

Cheers, and Merry Christmas to all,

Fred Monson

P.S. Maximow, Bloom, Ham and Fawcett. Those from whose hard-bound books I learned, and then, taught this stuff.

Frederick C Monson, PhD
Center for Microanalysis and Imaging, Research and Training (CMIRT) West Chester University of Pannsylvania West Chester, PA, 1938
610-738-0437
fmonson-at-wcupa.edu
________________________________________
X-from: PhillipsT-at-missouri.edu [PhillipsT-at-missouri.edu]
Sent: Thursday, December 06, 2012 6:55 PM
To: Monson, Frederick

Having worked on mucosal tissues most of my professional career, I think I know the answer to my own question but I can't prove it!

I have always used the phrase "lamina propria" to refer to the connective tissue layer below the epithelium and above the muscularis mucosae. I have assumed the phrase "substantia propria" is for a similar layer but more correctly used for the connective tissue layer below an epithelium that doesn't have a muscularis mucosae and therefore no distinction between a lamina propria and underlying submucosa. But despite aggressive Googling, I am unable to find a definition of substantia propria from an authoritative source. Is this something I made up in my head (perhaps they are true synonyms that reflect country of use) or is my definition precise? Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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I appreciate the thoughts but not remained less than fully satisfied. The substantia propria is also used to describe connective tissue in the conjunctiva and sclera, in the ear in relation to the membrane tympani and less commonly in the uterus. The citation from Gray's on cornea might indeed be an early "defining moment" that entrenched the use of this term for mostly ocular tissues. My angst over using it results mostly from not understanding why a novel term is used here. Was there a unique feature (lamellated fibers?) that makes it a substantia propria or simply a historical practice for those in the ocular milieu. I can't speak to Harvard's practice on teaching gross anatomy but when I played a small part in teaching Histology there, the course was legendary for the quality of job done by Marian Neutra and Dan Goodenough. Sadly, as in many Histology classes, ocular tissues weren't discussed.

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
X-from: FMonson-at-wcupa.edu [mailto:FMonson-at-wcupa.edu]
Sent: Thursday, December 06, 2012 7:39 PM
To: Phillips, Thomas E.

http://education.yahoo.com/reference/gray/subjects/subject/225 [cornea]

http://education.yahoo.com/reference/gray/illustrations/figure?id=871

http://medical-dictionary.thefreedictionary.com/Substantia+propria [synonyms at bottom of paragraph]

I remember the rumor, years ago, that Harvard Med, for a brief moment, declared 'gross' anatomy for medical students to be optional. Hmmmm!

I hear complaints from academia on high that there are no histologists to analyze research specimens. If there are none, then there can be no trainers, expecially if the 'specialty' has been labelled archaic. As I peruse the current hardbacks in histology, they are found to be directed to pathologists. Histology has become relegated to soft cover bindings.

Nomenclature is a pain. Thus, there are those who spend their careers relegating old terminology to oblivion by not learning it in the first place. In this case, it turns out that 'substantia propria' was used to describe poorly fixed, compact material in the cornea that lay between layers #2 and #4 of 5 of Grey's Anatomy, Fig. 871 [cited above].

It is not lamina propria, and it would be appropriate to print this response, file it, and spend a few evenings at the local pub trying to target that part of the neural net that holds the term 'substantia'. Thus, will 'propria' be retained unencumbered with an archaic Latin term whose meaning in Emglish is, at best, vague - at least according to 1 or 2 of the last Latin experts in the world.

I hope all of this helps, because you and I, Prof Phillips, will likely be the last of our kind to discuss this part of histologic anatomy.

Cheers, and Merry Christmas to all,

Fred Monson

P.S. Maximow, Bloom, Ham and Fawcett. Those from whose hard-bound books I learned, and then, taught this stuff.

Frederick C Monson, PhD
Center for Microanalysis and Imaging, Research and Training (CMIRT) West Chester University of Pannsylvania West Chester, PA, 1938
610-738-0437
fmonson-at-wcupa.edu
________________________________________
X-from: PhillipsT-at-missouri.edu [PhillipsT-at-missouri.edu]
Sent: Thursday, December 06, 2012 6:55 PM
To: Monson, Frederick

Having worked on mucosal tissues most of my professional career, I think I know the answer to my own question but I can't prove it!

I have always used the phrase "lamina propria" to refer to the connective tissue layer below the epithelium and above the muscularis mucosae. I have assumed the phrase "substantia propria" is for a similar layer but more correctly used for the connective tissue layer below an epithelium that doesn't have a muscularis mucosae and therefore no distinction between a lamina propria and underlying submucosa. But despite aggressive Googling, I am unable to find a definition of substantia propria from an authoritative source. Is this something I made up in my head (perhaps they are true synonyms that reflect country of use) or is my definition precise? Thanks. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Try using a pin hole aperture in place of the ocular (assuming you don't have a focusing Bertand lens) to see the objective BFP. Just take the ocular out of the scope, wrap a little Al foil over the opening and make a pin hole in the center of the foil. This will let you see the BFP. It will be tiny, but visible.

If you have a ground glass between the filament and your sample, you will not be able to see the filament, but you may be able to focus the lamp condenser (not the substage condenser) so that the ground glass is in the objective BFP. The leaves of the substage condenser should also be in focus in the objective BFP. Please make sure they are open to the edge of the field, you'll adjust then later for the specimen contrast you want.

Close the lamp condenser iris down to a pin point and focus that as sharply, with the sub stage condenser, as possible in the plane of the sample. Then open it so the leaves are just out side the field of view.

You may see color fringes, blue and red, depending on the focus. I pick a color in between - purple.

It sounds like you have a flip in lens on your condenser. Use the lower NA setting (flip in out) for lower NA objectives. The 1.25 NA is for oil or water (I've read about glycerin immersion but never seen one) immersions or very high dry objectives.

Oh--Start with the scope focused on some object, I prefer a diatom, but that's just me.

If this doesn't work (my scope is an Leitz monocular petrographic with a mirror) e-mail me and I scan a couple pages out of a work book I learned on which may, will have, more clarity.

Good luck, we're all rooting for you.

Frank

-----Original Message-----
X-from: dcristofori-at-unive.it [mailto:dcristofori-at-unive.it]
Sent: Friday, December 07, 2012 7:09 AM
To: Frank Karl

Dear Listers,
I'm quite embarassed asking your help just for setting up Koehler
condition in my Leica DM LM. I studied it a long time ago but I've never
needed to use it. Now it's time, but I cannot manage to get the right
condition.

With the specimen in focus, field aperture rim appears quite blurred
becuase of chromatic aberration, but maybe it's normal given the
condenser is just an achromat (CL/PH 0.90/1.25 S1); and it looks almost
in focus only if condenser aperture is almost closed and the condenser
is at the end of its focus range, by the sample side.
Besides I'm not able to find the filament image in objective bfp,
watching in the tube without eyepieces while moving the condenser along
its whole focus range. Maybe this image is just so demagnified that I
didn't realize its presence.

Could any one give me some very basic hint?
Thanks a lot.

Davide Cristofori

~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~
Davide Cristofori

direct phone = 041.234.67.26
e-mail = dcristofori[at]unive[dot]it

Universit� Ca' Foscari Venezia
Dipartimento di Sc. Molecolari e Nanosistemi
Via Torino, 155b I-30172 Mestre (VE) Italy
~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~

==============================Original Headers==============================
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This email and any of its attachments may contain confidential information intended only for the use of the addressee(s). If the reader of this email is not the intended recipient or the employee or agent responsible for delivering it to the intended recipient, you are hereby notified that any dissemination or copying of this email is strictly prohibited. If you have received this email in error, please notify us by return email at info-at-ardl.com, permanently delete the email, and destroy any printouts. If this email contains test data and/or draft reports, you are hereby notified that only a signed original test report will contain official results, a copy of which resides in the project folder located at ARDL, Inc. Thank you. Akron Rubber Development Laboratory, Inc.

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As my way of saying thanks to Fred, Wolfgang and Geoff for their helpful comments, I share with the list this link to an older story I happened on today about the world's smallest snowman - built with the assistance of a FIB.� You will need to cut and paste the link since I believe Nestor's filter will block me pasting a direct link.

http://www.telegraph.co.uk/news/newstopics/howaboutthat/6724969/Scientists-create-the-worlds-smallest-snowman.html

Happy holidays, Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

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Hello All,

We were hoping for some help on a search we are working on. Perhaps, this opportunity might be right for you or someone you know, who could benefit from speaking with us.

We are executing a search for a person who has experience in the following key areas: Microscopy, TEM, technical knowledge of Asbestos, Geology, Mineralogy and other duties related to Asbestos analysis.

We are a confidential search firm with an excellent reputation for such and treat all inquiries accordingly. I can be reached at:

paul at jpamri.com or 866-712-1810 Paul Palazzolo, Senior Managing Partner, JPA

We thank you in advance for your time and consideration

==============================Original Headers==============================
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Thank you very much to everybody who replied with instructions,
documentation, suggestions, rootings and willingness to continue
supporting me. You are so many, and all fantastic!

From what you wrote, I think the condition I had to accept as the least
bad one was actually the Koehler - or quite close. Nevertheless, I was
definitely puzzled, and words by experienced people were just
fundamental to me.
Actually, I haven't yet checked for element alignments and positions, so
maybe there's still something to do. But now I've got plenty of
documentation to learn from, and I'm sure I'll find out if I have to fix
something else.
Thanks again!

Davide

~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~
Davide Cristofori

direct phone = 041.234.67.26
e-mail = dcristofori[at]unive[dot]it

Universit� Ca' Foscari Venezia
Dipartimento di Sc. Molecolari e Nanosistemi
Via Torino, 155b I-30172 Mestre (VE) Italy
~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~~~~~~

==============================Original Headers==============================
6, 18 -- From dcristofori-at-unive.it Fri Dec 7 14:34:38 2012
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Greetings from FSU!

The Florida State University Office of Research is organizing a two-day
symposium to celebrate the installation of the state-of-the-art probe
aberration corrected JEM-ARM200cF Transmission Electron Microscope. This
event will take place Wednesday, February 20, and Thursday, February 21,
2013 at the National High Magnetic Field Laboratory, Florida State
University, Tallahassee, Florida.

The first day includes the presentations by invited speakers and a general
overview of the microscopy facility. The second day is dedicated to live
demos by JEOL, Inc., Nanofactory Instruments Inc., and Protochips, Inc.
for in situ TEM holders.

The symposium details and registration information can be found on our
website:

http://www.research.fsu.edu/tem/meeting/program.html

Registration is required. The deadline for registering is February 11,
2013.

We hope you can join us for this symposium and look forward to seeing you
in sunny Florida this February!

Web site http://www.research.fsu.edu/tem/index.html

Yan Xin
Associate Scientist
National High Magnetic Field Laboratory
Florida State University
1800 E. Paul Dirac Drive
Tallahassee, FL 32310

Tel: 850 644 1529
Fax: 850 644 0867
e-mail: xin-at-magnet.fsu.edu

==============================Original Headers==============================
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Dear Colleagues,
�
Good time for a discussion around water cooling (temperature is under 0�C outside).
We have started our TEM (FEI G20) some 8 years ago, the cooling loop is completely isolated from light and it was filled with tap water and an anticontaminant called thermoclean. The temperature is set on 17�C but we had some periods of total inactivity so the water come to room temperature for several monthes. Since then I never changed the water, just added some more when needed. I did a microbiological test every year and it was always negative, until this year (minimal contamination, but still something).
I wondered if there is a standard accepted procedure for the water cooling: is there a need to change the whole water after some time?
Do you use an anticontaminant and which one? (and how)
�
I also have a colleague who is running another machine with a water cooling set on 38�C, which would be perfect to use as a bioreactor but he does not agree :-)
He tried tap water with thermoclean but he got a strong contamination after 2 monthes. I must say that the thermoclean is now 8 years old and although it is still young for a whisky�it may be too old and�it lost its activity so I advised my colleague�to buy�a new one. Anyway I would still be grateful if someone could give me a hint for a product which inhibits microbial growth at 38�C in a water cooling loop.
�
Please no answer from Australia or New-Zealand saying that they are enjoying 28�C on the beach (please be kind with my depression)
�
Thank you in advance

Stephane

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Please share - priority registration closes Dec 20, 2012

Advanced Microscopy LIVE Workshop
January 14-18, 2013
University of California, Santa Barbara

Offered by the Neuroscience Research Institute (NRI) and Department of
Molecular, Cellular, and Developmental Biology (MCDB).
See: https://www.lifesci.ucsb.edu/mcdb/events/imaging-live/

This workshop is five days of seminars and hands on instruction. Through
the course, participants will gain theoretical and practical experience
with light microscopy on live specimens. Lectures will cover basic to
advanced microscopy principals with an emphasis on live imaging
applications.

--
Mary Raven
Microscopy Facility Director
Neuroscience Research Institute &
Molecular, Cellular & Developmental Biology
Biology 2 Building, room 5173
The University of California
Santa Barbara, CA 93106-5060

2013Advanced Microscopy - LIVE Workshop
https://www.lifesci.ucsb.edu/mcdb/events/imaging-live/
Jan 14- Jan 18, 2013

Winter Break 2012/13 out Dec 21- back Jan 3

http://www.lifesci.ucsb.edu/~m_raven/
Phone: (805) 893 8702
Fax: (805) 893 2005

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Email: joseph.uknalis-at-ars.usda.gov
Name: Joe Uknalis

Organization: USDA

Title-Subject: [Filtered] regulator on CO2 tank - CPD

Message: Hi Folks-
I'd like to put a regulator on the Denton critical point dryer CO2 tank so I know how much I have
left before I start a run.

There seem to be plenty of dual regulators with the low side } 120psi output, but if you go to dual
with high pressure on both dials they become expensive:

http://store.cyberweld.com/smhiprre820s.html

Can I run this CPD with the lower pressure output?

Thanks

Joe
USDA-ERRC

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Email: rwchin-at-stanford.edu
Name: Richard Chin

Organization: Stanford University

Title-Subject: [Filtered] Position available: SEM & XPS -at- the Stanford Nanocharacterization Lab

Message: Nanocharacterization Science & Engineering Associate
Stanford Nanocharacterization Laboratory

General Description

The science and technology of the nanoscale is one of the most promising areas in science and
engineering today. The Stanford Nanocharacterization Laboratory (SNL) provide some of the most
advanced nano characterization equipment available. The instruments are available to all qualified
users in the Stanford community and to Stanford collaborators both locally and globally.

Working closely with faculty, staff and students, the Nanocharacterization Science & Engineering
Associate at the SNL will be part of a team to participate in scientific and engineering research
programs utilizing the instrumentation and capabilities available at SNL. The candidate provides key
support and guidance to researchers in nanoscale science and engineering.

The NSEA is responsible for operation, training, and helping to manage complex research equipment in
the SNL, specifically the dual-beam focused ion beam/scanning electron microscopes (FIB/SEM) and
X-ray photoelectron spectrometers (XPS), in coordination with existing staff, and to ensure that
equipment is used correctly and safely.

Responsibilities

Support research programs by giving researchers advice on best-practices on available research
instrumentation and capabilities
Coordinate and assist in installing and maintaining complex laboratory equipment
Oversee equipment operations including scheduled and unplanned maintenance
Schedule training and train researchers on the equipment
Establish and maintain training and operating procedures
Work closely with other researchers on their processes
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Provide guidance to others in the use of the equipment
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capabilities available at the SNL
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Collaborate with other staff to continuously seek improvements to support researcher needs
Support researchers needing in-depth analysis and assistance on a service-basis as needed
Assist in assuring compliance with Health & Safety regulations
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FIB/SEM and XPS techniques
Assist Property Administrator in the proper identification, tagging, and disposal of capital equipment
Oversee student helpers
Develop and maintain a personal training program in order to stay abreast of best practices on all
relevant equipment

Qualifications

Extensive practical experience in the operation and use of dual-beam FIB/SEM and XPS is required.
Experience in the daily operation and maintenance of FIB/SEMs and XPSs and auxiliary equipment
is required.
Experience in applying scientific and engineering principles and practices in advancing nanoscale
science and engineering research projects.
Discretion and judgment in work that is primarily intellectual and non-routine in order to help
graduate students use and interpret the results of advanced scientific instruments.
Experience in conducting independent research, writing scientific papers, and presenting
conference papers.
Good communication skills are required to interface with faculty, staff and students in their
research projects.
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guides is required.
Ability to act as a consultant about the tools and capabilities to the research community.
Ability to supervise student helpers and delegate appropriate tasks to others.
Masters degree in Materials Science or related discipline is required.
Two years of job-related experience is required.

A background check will be required for all final candidates.

Stanford University is an affirmative action, equal opportunity employer.

Thank you for your interest!

http://recruit.trovix.com/jobhostmaster/jobhost/ViewJobPostDetails.do?title=NANOCHARACTERIZATION+SCIENCE+AND+ENGINEERING+ASSOCIATE&jobPostId=uqw46qc57faizp42vuc2rbxjbg&accountId=de85ad313f8598db1c42b567a3df24a00497ba22&button=&action=viewDetails&tid=0207-kyckbavk6nfm7jvk7jbmclv7bg

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Sorry Joe,

You need the full cylinder pressure, 800psi, to have liquid CO2 for you CPD. The only way I know of estimating when the tank is going to run out of Liquid CO2 is by weighing the cylinder. The pressure inside the tank does not go down when you are using the liquid it will only go down when you have used all the liquid and only have CO2 gas in the cylinder.

Regards
John

John V Nailon
Operations Manager
Centre for Microscopy and Microanalysis
AIBN Building (75)
University of Queensland
St.Lucia QLD 4072
Phone: 07 3346 3988
Fax: 07 3346 3993
email: J.Nailon-at-uq.edu.au

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Email: joseph.uknalis-at-ars.usda.gov
Name: Joe Uknalis

Organization: USDA

Title-Subject: [Filtered] regulator on CO2 tank - CPD

Message: Hi Folks-
I'd like to put a regulator on the Denton critical point dryer CO2 tank so I know how much I have left before I start a run.

There seem to be plenty of dual regulators with the low side } 120psi output, but if you go to dual with high pressure on both dials they become expensive:

http://store.cyberweld.com/smhiprre820s.html

Can I run this CPD with the lower pressure output?

Thanks

Joe
USDA-ERRC

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On Dec 7, 2012, at 12:08 PM, PhillipsT-at-missouri.edu wrote:

} As my way of saying thanks to Fred, Wolfgang and Geoff for their
} helpful comments, I share with the list this link to an older story
} I happened on today about the world's smallest snowman - built with
} the assistance of a FIB. You will need to cut and paste the link
} since I believe Nestor's filter will block me pasting a direct link.

Dear Tom,
Having been caught in Nestor's filter myself, I was surprised that I
could just click on the link. I think the Mac Mail program
reactivated it. Anyhow, the platinum-and-styrofoam snowman was cool,
but can anyone make one out of amorphous ice? I think it can be done,
but does anyone have that much free time?
Yours,
Bill

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Good morning, good evening,
Dear Listers,

This message just to say thank you to all of the Listers, especially also to Prof. Phillips for his recent "Thanks"-Mailing on the subject {Histology question on lamina propria vs substantia}

as well as to say that I never had a problem with pasted direct www.links embedded in the message body of a Text-format Mail (given the link was written properly, hopefully see below: I have added again the link which was pasted by Prof. Phillips).
PC-System: MS Windows XP, MS Outlook.

Thanks for great answers, and excellent collegial help over the year,

with my best wishes to you all for a Merry Christmas,
a Happy, healthy, prosperous and successful New Year
/ Season's Greetings

Wolfgang MUSS PhD
EM-Lab
Pathology, SALK-LKH & PMU
General Hospital and Paracelsus Med. Univ.
SALZBURG-AUSTRIA
Secretary of SCUR
http://www.scur.org (this link typed, no RETURN: URL resembles only {text without a link function} = NOT {blue and underlined} ....] guessing it will show up underlined too when you receive the message.
http://www.scur.org
(Last link typed, RETURN: URL resembles even in Mail-TEXT-Format the usual hyperlink format {text with a link function} = {blue and underlined} ....]

wtivol-at-sbcglobal.net

} -----Urspr�ngliche Nachricht-----
} Von: wtivol-at-sbcglobal.net [mailto:wtivol-at-sbcglobal.net]
} Gesendet: Mittwoch, 12. Dezember 2012 04:56
} An: Mu� Wolfgang
} Betreff: [Microscopy] Re: Thanks
} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America
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}
} On Dec 7, 2012, at 12:08 PM, PhillipsT-at-missouri.edu wrote:
}
} } As my way of saying thanks to Fred, Wolfgang and Geoff for their
} } helpful comments, I share with the list this link to an older story
} } I happened on today about the world's smallest snowman - built with
} } the assistance of a FIB. You will need to cut and paste the link
} } since I believe Nestor's filter will block me pasting a direct link.

http://www.telegraph.co.uk/news/newstopics/howaboutthat/6724969/Scientists-create-the-worlds-smallest-snowman.html
[Link again added by W. Muss: here/originally the URL resembles only {text without a link function} = NOT {blue and underlined} ....]

} Dear Tom,
} Having been caught in Nestor's filter myself, I was surprised that I
} could just click on the link. I think the Mac Mail program
} reactivated it. Anyhow, the platinum-and-styrofoam snowman was cool,
} but can anyone make one out of amorphous ice? I think it can be done,
} but does anyone have that much free time?
} Yours,
} Bill
}
}
}
}
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13, 39 -- From W.Muss-at-salk.at Wed Dec 12 01:32:43 2012
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Joe,

I wouldn't bother. You're using a siphon CO2 tank, yes? So you're
withdrawing liquid CO2 from the tank, which means you won't see any
pressure drop until you've run out of liquid CO2, in which case it's
too late. The best way to know how much CO2 you have left is to have
the tank on a scale. The supply company should put in a specific
weight of CO2, usually 65 pounds, and there is (supposed to be) a
tare weight on the tank. When you've used say 64 pounds, change tanks.
If you're not using a siphon CO2, change tanks and get one. Food
grade is best (less contamination like oils).

Phil

} Email: joseph.uknalis-at-ars.usda.gov
} Name: Joe Uknalis
}
} Organization: USDA
}
} Title-Subject: [Filtered] regulator on CO2 tank - CPD
}
} Message: Hi Folks-
} I'd like to put a regulator on the Denton critical point dryer
} CO2 tank so I know how much I have
} left before I start a run.
}
} There seem to be plenty of dual regulators with the low side } 120psi
} output, but if you go to dual
} with high pressure on both dials they become expensive:
}
} http://store.cyberweld.com/smhiprre820s.html
}
}
} Can I run this CPD with the lower pressure output?
}
} Thanks
}
} Joe
} USDA-ERRC

--
Philip Oshel
Microscopy Facility Supervisor
Biology Department
024C Brooks Hall
Central Michigan University
Mt. Pleasant, MI 48859
(989) 774-3576

==============================Original Headers==============================
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Dear Listers,
Any technical know-how users of Aspex PSEM3025 out there?. I am having a problem of not having the vacuum ready signal on the control panel. This is the first pre-requisite to start the emitter. After being put to pump, even after a lenghty wait, the vacuum interlock panel window is not showing "Ready". The vac. reading is struck at 0.0032Torr. The section which shows "sample pressure" is reaching "high vac" shortly. I cleaned and regreased the specimen chamber interlock "O" ring, with no improvement.

Your advise and suggestions please.

Mohammed Yousuf Ph.D.
Senior Research Associate
Research and Graduate Studies (RGS)
Material Characterization Lab (152)
Texas A&M University in Qatar
Education City, PO. Box. 23874
Doha, State of Qatar.

Work: 00974-44230592
Mobile: 00974-55720907
mdyousuf-at-qu.edu.qa
mohammed.yousuf-at-qatar.tamu.edu
________________________________

رؤيتنا: أن تصبح جامعة قطر نموذجا للجامعة الوطنية في المنطقة، تتميز بنوعية التعليم والأبحاث، وبدورها الرائد في التنمية الاقتصادية والاجتماعية.

Our Vision: Qatar University shall be a model national university in the region, recognized for high quality education and research, and for being a leader of economic and social development.

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Dear Collective,

Just wondering if anyone has ever come up with a decent method for TEM imaging of starch inclusions in plant tissue (like soybean!) without having the field full of holes where starches saw their opportunity to escape. I expect not, but hope blooms eternal....

Happy Holidays to all!

Cheers,
Randy

Randy Tindall
Senior Research Specialist
Electron Microscopy Core Facility
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65201
(573) 884-5383

We Do Small Well!

www.emc.missouri.edu (dayjob)
http://nadiasyard.com (blog)
http://www.facebook.com/RandyTindallGalleries (photography)

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Dear Stephane,

(And others that may be interested...)

This from the FEI Technical Service Group:

We advise to use thermoclean DC, Please contact the supplier at supportAbioanalytic.de for more information if interested.

In reading your question I would at least add new thermoclean DC every year. I had enclosed two PDF's but the listserver stripped those. For Titan we can use the MSB procedure; for Tecnai we do not know yet but if it is a Tecnai with single chiller (I.E. without UT or Lorentz lens coils which can be recognized by having an additional cooler) this can also be used. In the document it is mentioned that it can be used up to 85 degrees but please contact the supplier for the dose and how often this needs to be replaced.

Hope you find this information useful. I should add that actually (other than the comments you make about people currently being on the beach etc.) there is a real variation worldwide regarding what the best treatment is depending on local water quality, acidity, cleanliness, additives etc. so you will find that different labs have different procedures. FEI tried to standardize the treatment protocol, but because of variations in water, acidity particularly, this is difficult to do.

Sincerely, Jan

} -----Original Message-----
} From: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: Monday, December 10, 2012 9:29 AM
} To: Ringnalda, Jan
} Subject: [Microscopy] Water cooling and contamination
}

} ------
}
} Dear Colleagues,
}
} Good time for a discussion around water cooling (temperature is under 0�C outside).
} We have started our TEM (FEI G20) some 8 years ago, the cooling loop is completely isolated from light and it was filled with tap water and an anticontaminant called thermoclean. The temperature is set on 17�C but we had some periods of total inactivity so the water come to room temperature for several monthes. Since then I never changed the water, just added some more when needed. I did a microbiological test every year and it was always negative, until this year (minimal contamination, but still something).
} I wondered if there is a standard accepted procedure for the water cooling: is there a need to change the whole water after some time?
} Do you use an anticontaminant and which one? (and how)
}
} I also have a colleague who is running another machine with a water cooling set on 38�C, which would be perfect to use as a bioreactor but he does not agree :-) He tried tap water with thermoclean but he got a strong contamination after 2 monthes. I must say that the thermoclean is now 8 years old and although it is still young for a whisky it may be too old and it lost its activity so I advised my colleague to buy a new one. Anyway I would still be grateful if someone could give me a hint for a product which inhibits microbial growth at 38�C in a water cooling loop.
}
} Please no answer from Australia or New-Zealand saying that they are
} enjoying 28�C on the beach (please be kind with my depression)
}
} Thank you in advance
}
} Stephane
}

==============================Original Headers==============================
14, 30 -- From prvs=3694D161A7=Jan.Ringnalda-at-fei.com Thu Dec 13 08:45:29 2012
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I'm new to this lab and the recent PM we had identified we had cooling water contamination and leaks in the FEI Technai. The service engineer said it is up to the lab to maintain the cooling system to the point of changing the water on a regular basis. Within the EM itself it is FEI's problem. So now we are instituting an annual schedule for that.

Tim Morken
Supervisor, Electron Microscopy/Neuromuscular Special Studies
Department of Pathology
UC San Francisco Medical Center

-----Original Message-----
X-from: Jan.Ringnalda-at-fei.com [mailto:Jan.Ringnalda-at-fei.com]
Sent: Thursday, December 13, 2012 6:54 AM
To: Morken, Timothy

Dear Stephane,

(And others that may be interested...)

This from the FEI Technical Service Group:

We advise to use thermoclean DC, Please contact the supplier at supportAbioanalytic.de for more information if interested.

In reading your question I would at least add new thermoclean DC every year. I had enclosed two PDF's but the listserver stripped those. For Titan we can use the MSB procedure; for Tecnai we do not know yet but if it is a Tecnai with single chiller (I.E. without UT or Lorentz lens coils which can be recognized by having an additional cooler) this can also be used. In the document it is mentioned that it can be used up to 85 degrees but please contact the supplier for the dose and how often this needs to be replaced.

Hope you find this information useful. I should add that actually (other than the comments you make about people currently being on the beach etc.) there is a real variation worldwide regarding what the best treatment is depending on local water quality, acidity, cleanliness, additives etc. so you will find that different labs have different procedures. FEI tried to standardize the treatment protocol, but because of variations in water, acidity particularly, this is difficult to do.

Sincerely, Jan

} -----Original Message-----
} From: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Sent: Monday, December 10, 2012 9:29 AM
} To: Ringnalda, Jan
} Subject: [Microscopy] Water cooling and contamination
}

} ------
}
} Dear Colleagues,
}
} Good time for a discussion around water cooling (temperature is under 0�C outside).
} We have started our TEM (FEI G20) some 8 years ago, the cooling loop is completely isolated from light and it was filled with tap water and an anticontaminant called thermoclean. The temperature is set on 17�C but we had some periods of total inactivity so the water come to room temperature for several monthes. Since then I never changed the water, just added some more when needed. I did a microbiological test every year and it was always negative, until this year (minimal contamination, but still something).
} I wondered if there is a standard accepted procedure for the water cooling: is there a need to change the whole water after some time?
} Do you use an anticontaminant and which one? (and how)
}
} I also have a colleague who is running another machine with a water cooling set on 38�C, which would be perfect to use as a bioreactor but he does not agree :-) He tried tap water with thermoclean but he got a strong contamination after 2 monthes. I must say that the thermoclean is now 8 years old and although it is still young for a whisky it may be too old and it lost its activity so I advised my colleague to buy a new one. Anyway I would still be grateful if someone could give me a hint for a product which inhibits microbial growth at 38�C in a water cooling loop.
}
} Please no answer from Australia or New-Zealand saying that they are
} enjoying 28�C on the beach (please be kind with my depression)
}
} Thank you in advance
}
} Stephane
}

==============================Original Headers==============================
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All,

When I posted the link in the reply I mistyped the supplier contact info:

Please contact the supplier at support-at-bioanalytic.de for more information if interested.

Also here is a link to their website:

http://www.bioanalytic.de/waterbath-stabilizers.en.html

Jan

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Hi Everyone
We are planning a CSI Indianapolis for the Family Affair at M&M 2013 and I
need your help. Over the last few years we have put together a few CSI's
geared to the places we are in. I have never been to Indianapolis and I
need a senario for a crime we can solve, possibly involving cars and race
tracks.

The CSI part of the Family Affair is to give focus for the tour of the
Exhibit Hall. The first part of Family Affair will be to put the clues
together, to look especially at materials identification and how they
break. We usually provide clues which will involve some light microscopy
first. We should have some vendors willing to give us time with edx
systems. The major part of Family Affair is to have fun with microscopy.
Delegates who would like to see a workshop in progress please join us in
Indianapolis Wednesday afternoon, August 7, 2013

As you can see this is still a fun piece of work in progress. Let your
imaginations run!
Elaine Humphrey, Frauke Hogue and Stuart McKernan

Dr. Elaine C. Humphrey
STEHM Technologist and Lab Manager
Bob Wright Science Centre A015
Advanced Microscopy Facility
University of Victoria, Canada

Lab: 250-853-3968
cell: 250-886-2068
website: http://www.stehm.uvic.ca {http://www.stehm.uvic.ca/}

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I walked into a fairly new site with an older scope, which flooded me with memories of questions about ...well...here we go..

Let's see ..that..which we don't: (help me out OEMs and techies..)

1)What temperature ranges in each em component make a long or short term difference?

2) Can different types pumps transmit vibrations to lenses, DPs, etc.

3) What type of chemical characteristics effect the instrument lines,the pumps, the hoses(!),..?

4) Does the instrument(s) or the cooling system have flow, pressure and temperature monitors with feedback correction and auto-shutdown mechanisms? Other related controls needed?

5) I have seen several cooling pump systems fail due to clogged filters in the system that I was not aware of..and it didn't look like they were intended to be checked (even hidden in the system)..so they just... fail.

6,7,... How many other situations are overlooked because they are "just cooling systems"?!

I corrected and modified an SEM cooling system(changed pump design) once, after the 2nd pump failure failure, to find out that resolution/interference fluctuations dramatically disappeared. It occurred to me that this is more important an issue at every scope(high/TEM and low/dp-SEM res.)than we might have noticed.
jeff
____________________________________________________________
Veteran Home Loans
Apply for VA Loans with competitive interest rates at Military.com.
http://thirdpartyoffers.juno.com/TGL3131/50ca727e8c645727e2aacst02vuc

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Hi Randy,

Starch is difficult to embed, no doubt about it. My best results are by embedding in Spurr's resin (infiltrate slowly�).

Good cheers for the holidays!

Howard

Howard Berg
Director, Integrated Microscopy Facility
Danforth Plant Science Center
975 North Warson Road
St. Louis, MO 63132

314-587-1261

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} Dear Collective,
}
} Just wondering if anyone has ever come up with a decent method for TEM imaging of starch inclusions in plant tissue (like soybean!) without having the field full of holes where starches saw their opportunity to escape. I expect not, but hope blooms eternal....
}
} Happy Holidays to all!
}
} Cheers,
} Randy
}
} Randy Tindall
} Senior Research Specialist
} Electron Microscopy Core Facility
} W125 Veterinary Medicine
} University of Missouri
} Columbia, MO 65201
} (573) 884-5383
}
} We Do Small Well!
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9, 31 -- Subject: Re: [Microscopy] TEM: Starch inclusions
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Email: jasantanderm-at-asu.edu
Name: Javier Santander

Organization: Arizona State University Biodesign Institute

Title-Subject: [Filtered] statistics for bacteria in TEM analysis

Message: We are conducting a drug-treatment study of gram-positive bacteria that will include TEM,
and part of the data analysis will involve doing percentage counts of bacteria cells that display
the effect of treatment. We are looking for a method of performing a statistical analysis that will
ensure validity in the numbers. Any information or references would be appreciated. Thank you-

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Email: xulingling723-at-gmail.com
Name: Lingling

Organization: Dalhousie university

Title-Subject: [Filtered] Artifact of a video

Message: Hello,

I asked the question before but I didn't give enough information about the problem. So I'm going to
explain it again.
I'm using a ZEISS AXIOVERT 200M INVERTED MICROSCOPE to record the process of a cylindrical fiber
being pulled
from a viscous solution. The process uses a syringe needle to perform the pulling so it is not
possible to use a cover-slide. As well, I use a 40X objective (for 400X final magnification) so I'm
sure that only an inverted microscope can be used because the liquid layer is about 1-2 mm thick.
There's always an artefact in the form of a black/white spot (either really black or very bright
depending on the focus)at the location where the viscous solution is lifted up by the fiber and
forms a "convex" structure. I'm not aware of anyway to use epi-illumination on an inverted
microscope. I do use phase contrast to visualize the fiber formation.
Does anyone know what caused the black/white spot?

Thanks in advance,

Lingling

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Hi Lingling,
Maybe someone more experienced in it can give a better explanation, but I
think that the black/white spot is caused exactly by that "convex" structure
you are observing. Its surface is at another angle, forms a type of lens on
its own, so it refracts the light differently and you see: (i) black because
the light refracted by the convex structure does not enter the objective; or
(ii) see white because the convex structure refracts the light in the focal
point of the objective.

That is my understanding from the point of my basic knowledge of optics.
Maybe someone else can explain it better, or correct me. And I am not sure
if that would be named an artifact...

Best,
J

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

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Name: Lingling

Organization: Dalhousie university

Title-Subject: [Filtered] Artifact of a video

Message: Hello,

I asked the question before but I didn't give enough information about the
problem. So I'm going to explain it again.
I'm using a ZEISS AXIOVERT 200M INVERTED MICROSCOPE to record the process of
a cylindrical fiber being pulled from a viscous solution. The process uses a
syringe needle to perform the pulling so it is not possible to use a
cover-slide. As well, I use a 40X objective (for 400X final magnification)
so I'm sure that only an inverted microscope can be used because the liquid
layer is about 1-2 mm thick.
There's always an artefact in the form of a black/white spot (either really
black or very bright depending on the focus)at the location where the
viscous solution is lifted up by the fiber and forms a "convex" structure.
I'm not aware of anyway to use epi-illumination on an inverted microscope. I
do use phase contrast to visualize the fiber formation.
Does anyone know what caused the black/white spot?

Thanks in advance,

Lingling

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Hi Mohammed.
I'm not familiar with the PSEM3025 but work on the earlier model PSEM's. I would suspect a problem somewhere with the vacuum system causing the turbo not to reach normal operating speed: the turbo pump (check if the blades are seized and make sure the vent valve is not open), turbo converter (check for power and the three LED's lit inside), check the fore line hose clamps are tight, and even check the oil level in the rough pump.

Hope this helps.

Happy Holidays,
Andy Zobel
Instrument Technician

R J Lee Group

www.rjlg.com

724.325.1776 Office
724.325.1776 Direct
724.733.1799 Fax

350 Hochberg Road | Monroeville, PA 15146

Please let us know if we've met your expectations on your project by visiting our Customer Survey.

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Sent: Wednesday, December 12, 2012 9:12 AM
To: Andy Zobel

Dear Listers,
Any technical know-how users of Aspex PSEM3025 out there?. I am having a problem of not having the vacuum ready signal on the control panel. This is the first pre-requisite to start the emitter. After being put to pump, even after a lenghty wait, the vacuum interlock panel window is not showing "Ready". The vac. reading is struck at 0.0032Torr. The section which shows "sample pressure" is reaching "high vac" shortly. I cleaned and regreased the specimen chamber interlock "O" ring, with no improvement.

Your advise and suggestions please.

Mohammed Yousuf Ph.D.
Senior Research Associate
Research and Graduate Studies (RGS)
Material Characterization Lab (152)
Texas A&M University in Qatar
Education City, PO. Box. 23874
Doha, State of Qatar.

Work: 00974-44230592
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mdyousuf-at-qu.edu.qa
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Our Vision: Qatar University shall be a model national university in the region, recognized for high quality education and research, and for being a leader of economic and social development.

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It's conceivable, if the polymer is clear, the cylinder is acting as a lens and creating your spot as a function of focus. If you're near the focal point I would expect to see a bright spot. Further away, the front surface of the object could be "missing" the refracted light rays and seeing a dark spot.

I'd try oblique illumination and see if you get the spot.

My 2 cents worth.

Stay safe

Frank

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Email: xulingling723-at-gmail.com
Name: Lingling

Organization: Dalhousie university

Title-Subject: [Filtered] Artifact of a video

Message: Hello,

I asked the question before but I didn't give enough information about the problem. So I'm going to
explain it again.
I'm using a ZEISS AXIOVERT 200M INVERTED MICROSCOPE to record the process of a cylindrical fiber
being pulled
from a viscous solution. The process uses a syringe needle to perform the pulling so it is not
possible to use a cover-slide. As well, I use a 40X objective (for 400X final magnification) so I'm
sure that only an inverted microscope can be used because the liquid layer is about 1-2 mm thick.
There's always an artefact in the form of a black/white spot (either really black or very bright
depending on the focus)at the location where the viscous solution is lifted up by the fiber and
forms a "convex" structure. I'm not aware of anyway to use epi-illumination on an inverted
microscope. I do use phase contrast to visualize the fiber formation.
Does anyone know what caused the black/white spot?

Thanks in advance,

Lingling

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Dear Listers,

I have a faculty member who would like to know if it is possible to embed an entire mouse brain in LR White and then section it. Has anyone out there tried this?

Tom Bargar
UNMC
Electron Microscopy Core Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu

==============================Original Headers==============================
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And how would you propose sectioning it? Your knives must be a lot wider than mine. I doubt you could cut it on a paraffin-style metal knife microtome. Tom

Thomas E. Phillips, Ph.D
Professor of Biological Sciences
Director, Molecular Cytology Core
2 Tucker Hall
University of Missouri
Columbia, MO 65211-7400
573-882-4712 (office)
573-882-0123 (fax)
phillipst-at-missouri.edu

http://www.biology.missouri.edu/faculty/phillips.html
http://www.biotech.missouri.edu/mcc/

-----Original Message-----
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To: Phillips, Thomas E.

Dear Listers,

I have a faculty member who would like to know if it is possible to embed an entire mouse brain in LR White and then section it. Has anyone out there tried this?

Tom Bargar
UNMC
Electron Microscopy Core Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu

==============================Original Headers==============================
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16, 31 -- From PhillipsT-at-missouri.edu Mon Dec 17 09:41:12 2012
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16, 31 -- Subject: RE: [Microscopy] Embedding in LR White
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From promotions_managerr-at-superposta.com Mon Dec 17 13:22:42 2012
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I did not but 0,35 sec googling gives this:
http://www.nature.com/nmeth/journal/v9/n12/full/nmeth.2213.html
http://www.ihcworld.com/smf/index.php?topic=2889.0
http://www.microscopy-analysis.com/news/researchers-prepare-whole-mouse-brain-sem
�
It seems that the faculty member will not be the first one.
�
Regards,
Stephane

X-from: "tbargar-at-unmc.edu" {tbargar-at-unmc.edu}
To: nizets2-at-yahoo.com
Sent: Monday, December 17, 2012 3:49 PM

Dear Listers,

I have a faculty member who would like to know if it is possible to embed an entire mouse brain in LR White and then section it.� Has anyone out there tried this?

Tom Bargar
UNMC
Electron Microscopy Core Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu

==============================Original Headers==============================
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Hi Lingling,
�
Since the phase contrast mechanism is based on differences in density, it is not very surprising that with a gel, a fiber and a syringe needle you get a strong contrast. Did you try to observe your material in brightfield mode? Hopefully you fiber is still visible in BF!
If your Axiovert 200M has fluorescence capability, you may try to label your fiber and your gels with 2 different colors.
�
Regards,
Stephane
�

X-from: "jmircheski-at-us.es" {jmircheski-at-us.es}
To: nizets2-at-yahoo.com
Sent: Friday, December 14, 2012 2:46 PM

Hi Lingling,
Maybe someone more experienced in it can give a better explanation, but I
think that the black/white spot is caused exactly by that "convex" structure
you are observing. Its surface is at another angle, forms a type of lens on
its own, so it refracts the light differently and you see: (i) black because
the light refracted by the convex structure does not enter the objective; or
(ii) see white because the convex structure refracts the light in the focal
point of the objective.

That is my understanding from the point of my basic knowledge of optics.
Maybe someone else can explain it better, or correct me. And I am not sure
if that would be named an artifact...

Best,
J

Dr.�Josif�Mircheski
____________________________________________________________________________
___
Instituto de Biomedicina de Sevilla (IBIS), Lab 108
Campus del Hospital Universitario Virgen del Rocio/CSIC/Univ. de Sevilla
Avda. Manuel Siurot S/N�
41013 Sevilla
Spain
�
Phone:+34-955923045
e-mail:�jmircheski-at-us.es
web: www.ibis-sevilla.es

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Email: xulingling723-at-gmail.com
Name: Lingling

Organization: Dalhousie university

Title-Subject: [Filtered] Artifact of a video

Message: Hello,

I asked the question before but I didn't give enough information about the
problem. So I'm going to explain it again.
I'm using a ZEISS AXIOVERT 200M INVERTED MICROSCOPE to record the process of
a cylindrical fiber being pulled from a viscous solution. The process uses a
syringe needle to perform the pulling so it is not possible to use a
cover-slide. As well, I use a 40X objective (for 400X final magnification)
so I'm sure that only an inverted microscope can be used because the liquid
layer is about 1-2 mm thick.
There's always an artefact in the form of a black/white spot (either really
black or very bright depending on the focus)at the location where the
viscous solution is lifted up by the fiber and forms a "convex" structure.
I'm not aware of anyway to use epi-illumination on an inverted microscope. I
do use phase contrast to visualize the fiber formation.
Does anyone know what caused the black/white spot?

Thanks in advance,

Lingling

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Follow-up to my message:
I had a look at the wonderful work of Prof Denk in Germany.
Embedding, imaging and cutting the whole brain are perhaps just the easiest steps!
Because if you analyse the sections by EM you'll end up with a huge amount of data.
If you read his paper:
http://ac.els-cdn.com/S0959438811002133/1-s2.0-S0959438811002133-main.pdf?_tid=c01a10c4-48fb-11e2-acb8-00000aab0f26&acdnat=1355825867_58990577c5690b1c13c0a396312cb658
you'll see that just 1 mm� of brain by EM gives 1 Petabyte of data!!
The work hours to analyse the data are best described in 10th of thousands.
In some dark corner of my memory I seem to remember that a group of scientists designed a computer game where the players has to delimit axons in EM pictures. This is a way to distribute the work hours around the world.

The person responsible for the analysis should better enjoy these christmas holidays because they may well be the last ones before long ;-)

Regards,
Stephane
�

----- Forwarded Message -----
X-from: Stephane Nizet {nizets2-at-yahoo.com}
To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
Cc:
Sent: Tuesday, December 18, 2012 11:08 AM

Dear Listers,

I have a faculty member who would like to know if it is possible to embed an entire mouse brain in LR White and then section it.� Has anyone out there tried this?

Tom Bargar
UNMC
Electron Microscopy Core Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu

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15, 50 -- Subject: Fw: [Microscopy] Embedding in LR White
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Hi,

I think it is not impossible. It can be done if you can break glass
knives that size. Diamond knives are limited to 8mm (cutting edge
length/histo knives), as far as I know. I myself used 10 mm glass
knives to dissect whole specimens of krill (Meganyctiphanes norvegica)
embedded in LR White. Semi thin sectioning as well as ultra thin
sectioning worked well, but glass knives are quite "soft", so I used
about 100 for an animal of about 5 cm length. And my sections where
quite small compared to a mouse brain (} 5x5 mm). If the sections get
too large, it is difficult to transfer them from the water bath to the
staining trays...

Have fun!

:-) Torsten

}
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} Dear Listers,
}
} I have a faculty member who would like to know if it is possible to
} embed an entire mouse brain in LR White and then section it.� Has
} anyone out there tried this?
}
} Tom Bargar
} UNMC
} Electron Microscopy Core Facility
} 986395 Nebraska Medical Center
} Omaha, NE 68198-6395
} 402-559-7347
} tbargar-at-unmc.edu
}
}
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Hi, Stephane,

You're right! A group of scientists from Harvard, MIT and Max Planck Institute put their heads together and developed a computer game, which launched on the public domain last week, called "Eye-Wire", that allows budding scientists to help trace the neural network of the nerves from the eye to the brain. Here's the link to their announcement:

http://blog.eyewire.org/eyewire-the-official-launch/?goback=.gde_80899_member_194622760

and the link to the game:

https://eyewire.org/

Ed

Edward Haller, Lab Manager
University of South Florida
Integrative Biology Department
Electron Microscopy Core
SCA 110
4202 East Fowler Avenue
Tampa, FL 33620
813-974-2676
ehaller-at-usf.edu
Office: ISA 1046
________________________________________
X-from: nizets2-at-yahoo.com [nizets2-at-yahoo.com]
Sent: Tuesday, December 18, 2012 5:31 AM
To: Haller, Edward

Follow-up to my message:
I had a look at the wonderful work of Prof Denk in Germany.
Embedding, imaging and cutting the whole brain are perhaps just the easiest steps!
Because if you analyse the sections by EM you'll end up with a huge amount of data.
If you read his paper:
http://ac.els-cdn.com/S0959438811002133/1-s2.0-S0959438811002133-main.pdf?_tid=c01a10c4-48fb-11e2-acb8-00000aab0f26&acdnat=1355825867_58990577c5690b1c13c0a396312cb658
you'll see that just 1 mm� of brain by EM gives 1 Petabyte of data!!
The work hours to analyse the data are best described in 10th of thousands.
In some dark corner of my memory I seem to remember that a group of scientists designed a computer game where the players has to delimit axons in EM pictures. This is a way to distribute the work hours around the world.

The person responsible for the analysis should better enjoy these christmas holidays because they may well be the last ones before long ;-)

Regards,
Stephane

----- Forwarded Message -----
X-from: Stephane Nizet {nizets2-at-yahoo.com}
To: "microscopy-at-microscopy.com" {microscopy-at-microscopy.com}
Cc:
Sent: Tuesday, December 18, 2012 11:08 AM

Dear Listers,

I have a faculty member who would like to know if it is possible to embed an entire mouse brain in LR White and then section it. Has anyone out there tried this?

Tom Bargar
UNMC
Electron Microscopy Core Facility
986395 Nebraska Medical Center
Omaha, NE 68198-6395
402-559-7347
tbargar-at-unmc.edu

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Name: Bobbie Schneider

Organization: Fred Hutch Cancer Research Center

Title-Subject: [Filtered] Carbon Evaporators

Message: Does anyone have any recommendations for a carbon evaporator. We would need to evaporate
metal and carbon. We also use it for glow discharging our grids.

Thank you,
Bobbie Schneider
FHCRC

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Email: bschneid-at-fhcrc.org
Name: Bobbie Schneider

Organization: Fred Hutch Cancer Research Center

Title-Subject: [Filtered] Carbon Evaporators

Message: Does anyone have any recommendations for a carbon evaporator. We would need to evaporate
metal and carbon. We also use it for glow discharging our grids.

Thank you,
Bobbie Schneider
FHCRC

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To start - I'm a neophyte in metalographic crystalography.

O.k., I have a user we're looking for grain structure size changes in stainless
steel 316. Samples are some wires embedded in phenoloic resin for
polishing. Their polished pretty well. Following a recommmendation we've
etched with a solution of 4g picric acid in 100ml ethanol. But we are not
seeing the grain structure in reflected light microscopy. (Moving to SEM later
today )

Is this a good etchant for SS 314? How long would you recommend etching
for?

Any other suggestions or recommendations?

Thank you one and all!

Richard E. Edelmann, Ph.D., Director
Center for Advanced Microscopy & Imaging
9C Upham Hall
Miami University, Oxford, OH 45056
Ph: 513.529.5712 Fax: 513.529.4243
E-mail: edelmare-at-muohio.edu
http://www.cami.muohio.edu

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Email: cdr32-at-duke.edu
Name: Catherine Reyes

Organization: Duke University

Title-Subject: [Filtered] Job: Research Scientist - Materials Characterization

Message: The Triangle Materials Research Science and Engineering Center (MRSEC) at Duke University
is seeking an individual with superb technical skills and a high level of expertise in experimental
aspects of characterization of colloidal materials and polymeric assemblies. The successful
candidate will be intimately familiar with at least one of the following techniques: transmission
electron microscopy (particularly cryo techniques) or advanced scattering methods (particularly
small angle X-ray or neutron scattering). Candidates with a combined background in cryo-TEM and
small angle scattering are preferred. The primary responsibilities of this individual will be data
acquisition using these techniques, data reduction and interpretation, and training of students and
post-doctoral fellows in these techniques. Other responsibilities include laboratory management and
instrument maintenance. The successful candidate must be comfortable taking initiative and fostering
a collaborative approach to working with a diverse group of faculty & students. Excellent
communication skills (verbal and written) are required. A bachelors degree or higher is preferred.
For more information, visit this site: http://mrsec.duke.edu/employment

This position will be based at Duke University in Durham, North Carolina.

Applications should contain a cover letter, complete curriculum vitae, and names and addresses of at
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Duke University is an equal opportunity institution. Duke is committed to recruiting, hiring, and
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Name: Judy King

Organization: West Virginia University - Pathology

Title-Subject: [Filtered] Prep for platelets for TEM

Message: Does anyone have a good procedure to prepare human platelets for TEM (type of tube, steps
to follow, etc.) ?

Judy King
jrkjack-at-aol.com

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Email: michael.cammer-at-med.nyu.edu
Name: Michael Cammer

Organization: Skirball Institute of Biomolecular Medicine

Title-Subject: [Filtered] looking for type K insert vendor

Message: We need one or two type K stage inserts that hold 96 well plates for our Ludl, Zeiss and
Nikon stages.

Would somebody please recommend vendors.

Thank you very much.
________________________________________________________
Michael Cammer, Assistant Research Scientist
Skirball Institute of Biomolecular Medicine
Lab: (212) 263-3208 Cell: (914) 309-3270

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Organization: NASA Johnson Space Center

Title-Subject: [Filtered] SEM Microscopist Openning at the NASA Johnson Space Center

Message: https://jacobsexternal-jacobstechnology.icims.com/jobs/11039/electron-microscopy-scientist/job

The Electron Microscopy Scientist will:

Maintain and operate two Scanning Electron Microscopes (SEM), a JEOL 5910LV and a JEOL 7600F.
(http://ares.jsc.nasa.gov/ares/kr_laboratories/SEM.cfm)
Coordinate all vendor service for the instruments.
Train and assist researchers and students in the use of the instruments.
Support peer-review research through high-quality analyses of astromaterials with the instruments.
Additional responsibilities may include development of new analytical techniques.
Perform other duties as required.

Qualifications:

Required Education/Experience/Skills:

A BS degree from an accredited university in an applicable geoscience discipline.
Experience in operating an SEM instrument.
Solid understanding of the theory and practice of electron microscopy analysis and data reduction.
Preferences:

Graduate degree(s) with experience in the field.
Knowledge of geological and planetary mineralogy is a plus.
Ability to interpret analyses in a geologically meaningful way.
Experience with a Focused Ion Beam (FIB) instrument and/or an electron microprobe is a plus.
Experience working effectively in interdisciplinary teams.
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What a time not to have my reference books. I'd recommend George Vander Voort's "Metallography: Principles and Practice." It is by far the best publication on the topics of etching metals. The photomicrographs are worth the price alone!

Etching stainless can be difficult, you may have to resort to electro etch. But in any case the better the final polish the better your results. I'd go to 1 micro diamond with a mirror finish. Absolutely no scratches to the eye and only a few tiny scratches under a magnifying glass. The etchants will accent any scratch or damage from under a scratch.

I'd first try,
Marbles (50ml concentrated HCl and 25 mls of saturated cuprous sulfate)
Or
Kallings (2g cupric chloride in 40 mls methanol 40mls water 40mls HCl)

While not always a sure fire reagent for stainless, but great on carbon steels try a mix of
50% nital and 50% picral

You can find other formulas for etchants on line as well as variations of the above formulas.

I'd etch lightly, check it under a stereomicroscope and then re-etch if I need to. If you over etch you can polish it out (generally speaking) by hand with your 1 micron diamond. Etching is as much an art as science. Good luck.

I would switch to methanol (it's not that poisonous) because mixing nitric acid (an oxidizer) with alcohols is dangerous. The smaller alcohols are more stable.

-----Original Message-----
X-from: edelmare-at-miamioh.edu [mailto:edelmare-at-miamioh.edu]
Sent: Tuesday, December 18, 2012 4:59 PM
To: Frank Karl

To start - I'm a neophyte in metalographic crystalography.

O.k., I have a user we're looking for grain structure size changes in stainless
steel 316. Samples are some wires embedded in phenoloic resin for
polishing. Their polished pretty well. Following a recommmendation we've
etched with a solution of 4g picric acid in 100ml ethanol. But we are not
seeing the grain structure in reflected light microscopy. (Moving to SEM later
today )

Is this a good etchant for SS 314? How long would you recommend etching
for?

Any other suggestions or recommendations?

Thank you one and all!

Richard E. Edelmann, Ph.D., Director
Center for Advanced Microscopy & Imaging
9C Upham Hall
Miami University, Oxford, OH 45056
Ph: 513.529.5712 Fax: 513.529.4243
E-mail: edelmare-at-muohio.edu
http://www.cami.muohio.edu

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I have tried a number of different etches including Kallings and
oxalic/electrolytic to reveal grain structure on severely "worked"
stainless (spring) wire. My wire was type 302 if memory serves. None
were very successful. Some structure was barely perceptible but nothing
like that obtained from wrought, recrystallized material. Etched both
lightly and to the point of pitting without revealing decent structure.
Apparently the problem has to do with the extremely deformed grains from
wire drawing. BSE grain channeling contrast was a little murky due the
the high strain levels but about as good as etching.
I would like to hear if you have any success.

Woody

On 12/18/2012 1:59 PM, edelmare-at-miamioh.edu wrote:
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} To start - I'm a neophyte in metalographic crystalography.
}
} O.k., I have a user we're looking for grain structure size changes in stainless
} steel 316. Samples are some wires embedded in phenoloic resin for
} polishing. Their polished pretty well. Following a recommmendation we've
} etched with a solution of 4g picric acid in 100ml ethanol. But we are not
} seeing the grain structure in reflected light microscopy. (Moving to SEM later
} today )
}
} Is this a good etchant for SS 314? How long would you recommend etching
} for?
}
} Any other suggestions or recommendations?
}
} Thank you one and all!
}
}
} Richard E. Edelmann, Ph.D., Director
} Center for Advanced Microscopy& Imaging
} 9C Upham Hall
} Miami University, Oxford, OH 45056
} Ph: 513.529.5712 Fax: 513.529.4243
} E-mail: edelmare-at-muohio.edu
} http://www.cami.muohio.edu
}
}

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Email: rfoley-at-uab.edu
Name: Robin Foley

Organization: UAB

Title-Subject: [Filtered] RE: Etching Stainless Steel 316

Message: Stainless is a pain! What makes it stainless is what makes it hard to etch....

Unless the grain size is very small - if you don't see the grains on the optical microscope, you
won't on the SEM. Also, if you haven't done much polished samples on the SEM -you'll find you need
to kick up the contrast much more than usual. There isn't much contrast (better optically!). As
long as the grain size is large enough, SEM is not required for grain size determination in
stainless. But for either equipment - it needs to be etched!

We do the following:

For polishing:
We usually finish polishing with Buehler MasterPrep after 1 micron diamond. It's expensive but
effective. It's 0.05 micron alumina in a proprietary solution. Wear gloves and safety glasses,
it's slightly corrosive. This polish is not required but I have found it is worth the money to
reduce times that my students spend struggling in the lab. Warning - for some alloys, it will
remove inclusions (like oxide inclusions) so if you are interested in the inclusions, use of the
Buehler MasterPrep may not be worthwhile.

For etching:
The best etch is generally electrolytic but I'm always willing to try something else first since I'm
not skilled at electrolytic. Simple picric in ethanol is an etch for carbon and low alloy steels
and won't work for stainless. There is a picric-HCL-ethanol etch that works for martensitic
stainless steels (called Villella's) but I've never found it to work for austenitics. The alloying
level is much higher in austenitics so they are a different world.

What I find almost always works on stainless (both cast and wrought) is one of the very, very nasty
"regia" etchants including (from George Vander Voorts Metallography Principles and Practice):

1) 1 part HNO3, 1 part HCl, 1 part water (General purpose etch for most stainless steels. Stir
solution during etching (20C) for uniform, stain free results. Outlines constituents, reveals grain
structure.)

2) 4 parts HCL, 3 parts HNO3, 4 parts water (Use same procedure as number 1).

3) 15 ml HCL, 5 Ml HNO3 (agua regia. For austenitic grades. Use fresh. Use at 20C for about 5s.
Attacks sigma, outlines carbides. After 20s, sigma compe,tely dissolved. Reveals grain
boundaries. DO NOT STORE ETCHANT.

4) 3 parts HCL, 1 part HNO3, 1 part glycerol (Glyceregia. For austenitic grades. Reveals grain
structure, outlines sigma and carbides).

WARNING! These etchants are nasty and you should use a fume hood with complete coverage with safety
gear. I do not store any of these etchants as the book states for some of the similar compositions
- Do not store. Many, many, years ago, as an undergraduate we had an etchant that was stored
explode so I've always paid attention whenever that is stated in a book.

Comments:
I put the etchant in a beaker and dip the sample in with tongs and (while wearing all the safety
gear!) gently move the sample around. Otherwise you get bubbles from the etchant that form and
leave unetched, perfect circles all over your sample. Very irritating.

Leave plenty of room in the beaker so that etchant does not splash out when you dip and stir. Add
enough etchant so the mounted sample is covered, even when you raise it slightly to gently stir.

It's better to have the sample face up so you can see it. It will become cloudy when it is etched.
This may be difficult to see if the diameter of the wire is small. Wait for it to get a bit
cloudy and then check it on the microscope. When etching is complete, you should see all the grain
boundaries. You don't want to overetch or the smaple becomes to three-dimensional for optical
microscopy. You can etch - check on scope - etch - check on scope, etc. You don't have to repolish
if you underetch. You do have to repolish if you overetch. Usually the last polishing step will
remove the overetch.

You do know that dry picric is explosive right? It's shock sensitive and quite dangerous. I used
it for years without knowing this and fortunately, never exploded it. Years ago, the safety people
came over for an inspection and found a bottle of picric that was dried out (it's shipped submerged
in water) and had a heart attack. They dressed in their bomb suits and submerged the entire bottle
in water. It was scary to think I had been using it for years with no knowledge of the danger. USE
YOUR MSDS!!! Also, if the picric fumes build up in a hood they can crytallize and become explosive.
We have a picric hood that washes down to help with this problem. In general, I avoid using the
picric unless necessary. I keep it (submerged in water) but only use it when I have to. Sorry for
the safety lecture but I am a worrier!!

Good luck and feel free to contact me off line with questions if you don't find this or the other
recommendations to work.

Robin Foley (old metallurgist that still etches many different cast and wrought stainless alloys on
a regular basis!)

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Title-Subject: [Filtered] Embedding Problem with the plastic -spurrs

Message: Holidays greetings to all!
We have been having problem with the spurrs imbedding: It always has a soft center area of the
tissue even the size of the tissue is not big. The softness is too bad to be cut even
thick(1wm)sections.
I am wondering if the problem come from the chemical of the set( always from EMS) since there are
few labs has the same problem here.
I appreciate any info from any body.
Gina Zhang

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Regarding the Explosivity of Picric Acid:

"You do know that dry picric is explosive right? It's shock sensitive and
quite dangerous. I used
it for years without knowing this and fortunately, never exploded it. Years
ago, the safety people
came over for an inspection and found a bottle of picric that was dried out
(it's shipped submerged
in water) and had a heart attack"

Per George Vander Voort:

"Fortunately, there have been no documented cases of explosions from picric
acid in laboratories, according to Phifer [1]. If it is wet with water, it
is not an explosive hazard and any attempt to blow it up by a bomb squad
will only result in picric acid being spread all over the immediate area.
The concern has always been in finding an old bottle that has dried up
producing dehydrated picric acid, and if it has a metal cap, rather than a
plastic cap. In such a case, shock-sensitive metallic picrates may have
formed at the cap-bottle interface. The solution is to have a robot pick it
up and re-hydrate the picric acid after opening the bottle under water. If
the cap is plastic, and the acid has dried out, friction from opening the
cap could cause detonation. The solution here is to place the bottle in a
large bucket or tank of water and allow water to dissolve any dried picric
acid on the cap threads. Leave the bottle in the water for a few days until
some water can be seen inside the bottle. Then, while under water, open the
lid and re-hydrate the picric acid.

Obviously, the wise lab manager checks the picric acid bottle periodically
(which can vary with lab usage of picric in etchants) to make sure that the
picric acid remains wet. Today, bottles are sold with at least 30% water
content. A good practice is to keep a log of when the bottle has been
checked and when water is added. Also, use only plastic or glass spatulas to
remove picric from the bottle and add it to the etchant. Do not use metal
spatulas and clean the cap and threads on the bottle and on the cap with a
wet paper towel. If you have copper piping, do not dispose of picric acid by
pouring it down the drain as explosive metallic salts could form."

My Opinion (having created some shock sensitive peroxides, and managed them
[and others'] as well):

If it's dry, keep it dry. Not a significant risk.
If it's wet, don't let it dry and saltate or react with metals. And again,
it will not be a significant risk.

I have both forms and they are kept that way dry/dry, wet/wet.
I have handled dry picric a lot over the years and never had an issue, as
long as it hadn't dehydrated from solution.

postscript - A fusion prep of Picric forms beautiful crystals (under
polarized light) that often show grain boundary migration.

Tony

.........................................
Andrew Anthony "Tony" Havics, CHMM, CIH, PE
Environmental, Health & Safety, Microscopy, & Forensic Engineering
pH2, LLC
5250 E US Highway 36, Suite 830
Avon, IN 46123
(317) 718-7020 Office
(317) 718-7038 Fax
(317) 409-3238 Cell
aahavics-at-pH2LLC.com
www.ph2LLC.com

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Name: Robin Foley

Organization: UAB

Title-Subject: [Filtered] RE: Etching Stainless Steel 316

Message: Stainless is a pain! What makes it stainless is what makes it hard
to etch....

Unless the grain size is very small - if you don't see the grains on the
optical microscope, you
won't on the SEM. Also, if you haven't done much polished samples on the
SEM -you'll find you need
to kick up the contrast much more than usual. There isn't much contrast
(better optically!). As
long as the grain size is large enough, SEM is not required for grain size
determination in
stainless. But for either equipment - it needs to be etched!

We do the following:

For polishing:
We usually finish polishing with Buehler MasterPrep after 1 micron diamond.
It's expensive but
effective. It's 0.05 micron alumina in a proprietary solution. Wear gloves
and safety glasses,
it's slightly corrosive. This polish is not required but I have found it is
worth the money to
reduce times that my students spend struggling in the lab. Warning - for
some alloys, it will
remove inclusions (like oxide inclusions) so if you are interested in the
inclusions, use of the
Buehler MasterPrep may not be worthwhile.

For etching:
The best etch is generally electrolytic but I'm always willing to try
something else first since I'm
not skilled at electrolytic. Simple picric in ethanol is an etch for carbon
and low alloy steels
and won't work for stainless. There is a picric-HCL-ethanol etch that works
for martensitic
stainless steels (called Villella's) but I've never found it to work for
austenitics. The alloying
level is much higher in austenitics so they are a different world.

What I find almost always works on stainless (both cast and wrought) is one
of the very, very nasty
"regia" etchants including (from George Vander Voorts Metallography
Principles and Practice):

1) 1 part HNO3, 1 part HCl, 1 part water (General purpose etch for most
stainless steels. Stir
solution during etching (20C) for uniform, stain free results. Outlines
constituents, reveals grain
structure.)

2) 4 parts HCL, 3 parts HNO3, 4 parts water (Use same procedure as number
1).

3) 15 ml HCL, 5 Ml HNO3 (agua regia. For austenitic grades. Use fresh.
Use at 20C for about 5s.
Attacks sigma, outlines carbides. After 20s, sigma compe,tely dissolved.
Reveals grain
boundaries. DO NOT STORE ETCHANT.

4) 3 parts HCL, 1 part HNO3, 1 part glycerol (Glyceregia. For austenitic
grades. Reveals grain
structure, outlines sigma and carbides).

WARNING! These etchants are nasty and you should use a fume hood with
complete coverage with safety
gear. I do not store any of these etchants as the book states for some of
the similar compositions
- Do not store. Many, many, years ago, as an undergraduate we had an
etchant that was stored
explode so I've always paid attention whenever that is stated in a book.

Comments:
I put the etchant in a beaker and dip the sample in with tongs and (while
wearing all the safety
gear!) gently move the sample around. Otherwise you get bubbles from the
etchant that form and
leave unetched, perfect circles all over your sample. Very irritating.

Leave plenty of room in the beaker so that etchant does not splash out when
you dip and stir. Add
enough etchant so the mounted sample is covered, even when you raise it
slightly to gently stir.

It's better to have the sample face up so you can see it. It will become
cloudy when it is etched.
This may be difficult to see if the diameter of the wire is small. Wait
for it to get a bit
cloudy and then check it on the microscope. When etching is complete, you
should see all the grain
boundaries. You don't want to overetch or the smaple becomes to
three-dimensional for optical
microscopy. You can etch - check on scope - etch - check on scope, etc.
You don't have to repolish
if you underetch. You do have to repolish if you overetch. Usually the
last polishing step will
remove the overetch.

You do know that dry picric is explosive right? It's shock sensitive and
quite dangerous. I used
it for years without knowing this and fortunately, never exploded it. Years
ago, the safety people
came over for an inspection and found a bottle of picric that was dried out
(it's shipped submerged
in water) and had a heart attack. They dressed in their bomb suits and
submerged the entire bottle
in water. It was scary to think I had been using it for years with no
knowledge of the danger. USE
YOUR MSDS!!! Also, if the picric fumes build up in a hood they can
crytallize and become explosive.
We have a picric hood that washes down to help with this problem. In
general, I avoid using the
picric unless necessary. I keep it (submerged in water) but only use it
when I have to. Sorry for
the safety lecture but I am a worrier!!

Good luck and feel free to contact me off line with questions if you don't
find this or the other
recommendations to work.

Robin Foley (old metallurgist that still etches many different cast and
wrought stainless alloys on
a regular basis!)

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Some excellent advice.

I worked with picric acid as an etchant component for almost three years. We stored excess picric acid solutions in all plastic bottles, no foil lined caps. We went through a lot of picric acid. Yes, they do ship it soaking water-logged wet, you only need to keep it damp. The dry material is sensitive. The salts of picric acid are much more sensitive to shock, especially the heavy metals like lead. But even the light salts, Na, K and NH3 are explosive. As one of my instructors said it's not much of an explosive, not even al Qaeda uses it.

Make sure you complete flush your drain and not leave any picric in the drain traps, especially lead elbows found in older buildings. If you use a lot of it, it can sublime and collect in hoods, so steam cleaning isn't a bad idea, but I never saw it done in our lab hood in the 2.9 years I was there. Maybe our volume wasn't high enough.

Picric acid stains protein so yellow fingers were our biggest worry.

I do remember a story from WW1 where a solder could be wounded by a pricate loaded shell, rescued under a truce flag dyed with picrate and them have his wounds treated with picrate at the hospital. Just part of my misspent youth,

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Sent: Thursday, December 20, 2012 10:42 AM
To: Frank Karl

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Email: rfoley-at-uab.edu
Name: Robin Foley

Organization: UAB

Title-Subject: [Filtered] RE: Etching Stainless Steel 316

Message: Stainless is a pain! What makes it stainless is what makes it hard to etch....

Unless the grain size is very small - if you don't see the grains on the optical microscope, you
won't on the SEM. Also, if you haven't done much polished samples on the SEM -you'll find you need
to kick up the contrast much more than usual. There isn't much contrast (better optically!). As
long as the grain size is large enough, SEM is not required for grain size determination in
stainless. But for either equipment - it needs to be etched!

We do the following:

For polishing:
We usually finish polishing with Buehler MasterPrep after 1 micron diamond. It's expensive but
effective. It's 0.05 micron alumina in a proprietary solution. Wear gloves and safety glasses,
it's slightly corrosive. This polish is not required but I have found it is worth the money to
reduce times that my students spend struggling in the lab. Warning - for some alloys, it will
remove inclusions (like oxide inclusions) so if you are interested in the inclusions, use of the
Buehler MasterPrep may not be worthwhile.

For etching:
The best etch is generally electrolytic but I'm always willing to try something else first since I'm
not skilled at electrolytic. Simple picric in ethanol is an etch for carbon and low alloy steels
and won't work for stainless. There is a picric-HCL-ethanol etch that works for martensitic
stainless steels (called Villella's) but I've never found it to work for austenitics. The alloying
level is much higher in austenitics so they are a different world.

What I find almost always works on stainless (both cast and wrought) is one of the very, very nasty
"regia" etchants including (from George Vander Voorts Metallography Principles and Practice):

1) 1 part HNO3, 1 part HCl, 1 part water (General purpose etch for most stainless steels. Stir
solution during etching (20C) for uniform, stain free results. Outlines constituents, reveals grain
structure.)

2) 4 parts HCL, 3 parts HNO3, 4 parts water (Use same procedure as number 1).

3) 15 ml HCL, 5 Ml HNO3 (agua regia. For austenitic grades. Use fresh. Use at 20C for about 5s.
Attacks sigma, outlines carbides. After 20s, sigma compe,tely dissolved. Reveals grain
boundaries. DO NOT STORE ETCHANT.

4) 3 parts HCL, 1 part HNO3, 1 part glycerol (Glyceregia. For austenitic grades. Reveals grain
structure, outlines sigma and carbides).

WARNING! These etchants are nasty and you should use a fume hood with complete coverage with safety
gear. I do not store any of these etchants as the book states for some of the similar compositions
- Do not store. Many, many, years ago, as an undergraduate we had an etchant that was stored
explode so I've always paid attention whenever that is stated in a book.

Comments:
I put the etchant in a beaker and dip the sample in with tongs and (while wearing all the safety
gear!) gently move the sample around. Otherwise you get bubbles from the etchant that form and
leave unetched, perfect circles all over your sample. Very irritating.

Leave plenty of room in the beaker so that etchant does not splash out when you dip and stir. Add
enough etchant so the mounted sample is covered, even when you raise it slightly to gently stir.

It's better to have the sample face up so you can see it. It will become cloudy when it is etched.
This may be difficult to see if the diameter of the wire is small. Wait for it to get a bit
cloudy and then check it on the microscope. When etching is complete, you should see all the grain
boundaries. You don't want to overetch or the smaple becomes to three-dimensional for optical
microscopy. You can etch - check on scope - etch - check on scope, etc. You don't have to repolish
if you underetch. You do have to repolish if you overetch. Usually the last polishing step will
remove the overetch.

You do know that dry picric is explosive right? It's shock sensitive and quite dangerous. I used
it for years without knowing this and fortunately, never exploded it. Years ago, the safety people
came over for an inspection and found a bottle of picric that was dried out (it's shipped submerged
in water) and had a heart attack. They dressed in their bomb suits and submerged the entire bottle
in water. It was scary to think I had been using it for years with no knowledge of the danger. USE
YOUR MSDS!!! Also, if the picric fumes build up in a hood they can crytallize and become explosive.
We have a picric hood that washes down to help with this problem. In general, I avoid using the
picric unless necessary. I keep it (submerged in water) but only use it when I have to. Sorry for
the safety lecture but I am a worrier!!

Good luck and feel free to contact me off line with questions if you don't find this or the other
recommendations to work.

Robin Foley (old metallurgist that still etches many different cast and wrought stainless alloys on
a regular basis!)

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Hi Gina,

My first thought was that your tissue is not completely dehydrated. How big are the tissue pieces? As I'm sure you know, ideally the pieces should be about a half-millimeter across for good infiltration at normal times. If they are bigger, you could extend the dehydration steps to include a couple more 100% acetone or propylene oxide steps, and then add a couple more infiltration steps and/or increase infiltration times.

Plant tissue is often harder to infiltrate than animal tissue. Adjust your schedules accordingly.

Good luck!

Merry Christmas to all!

Randy

Randy Tindall
Senior Research Specialist
Electron Microscopy Core Facility
W125 Veterinary Medicine
University of Missouri
Columbia, MO 65201
(573) 884-5383

We Do Small Well!

www.emc.missouri.edu (dayjob)
http://nadiasyard.com (blog)
http://www.facebook.com/RandyTindallGalleries (photography)

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Title-Subject: [Filtered] Embedding Problem with the plastic -spurrs

Message: Holidays greetings to all!
We have been having problem with the spurrs imbedding: It always has a soft center area of the tissue even the size of the tissue is not big. The softness is too bad to be cut even thick(1wm)sections.
I am wondering if the problem come from the chemical of the set( always from EMS) since there are few labs has the same problem here.
I appreciate any info from any body.
Gina Zhang

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31, 27 -- From tindallr-at-missouri.edu Thu Dec 20 10:31:41 2012
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I agree with Randy. It looks like you have a problem with resin infiltration (or may be with dehydration). I would suggest to increase incubation time (and number of changes) of resin/propylene oxide, resin, (and may be alcohol). For my tough specimens (usually bone) infiltration with resin can take up to 3 days. I would not suspect chemicals when seeing soft spots only in the center area of specimens.

Merry Christmas and Holiday Greetings!

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

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} We have been having problem with the spurrs imbedding: It always has a
} soft center area of the tissue even the size of the tissue is not big. The
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} I am wondering if the problem come from the chemical of the set( always
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Hi Gina,

Fully agree with Randy and Vladimir.

It is not very likely that there is anything the matter with the embedding reagents. Logic should tell us that if there is a heterogeneity pattern over the specimen, i.e. defined areas well embedded and sectioning well, other areas flawed, then it is a procedure/specimen issue.
When you say: "few labs has the same problem here" .. do they embed the same tissue as well? Have you tried the Spurr's from their lab which would seem the thing to do.

Have a nice Holiday Season, all!

Jan
i: www.aurion.nl

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} Email: gzhang-at-u.arizona.edu
} Name: Gina Zhang
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} Organization: U of A
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} Title-Subject: [Filtered] Embedding Problem with the plastic -spurrs
}
} Message: Holidays greetings to all!
} We have been having problem with the spurrs imbedding: It always has a soft center area of the tissue even the size of the tissue is not big. The softness is too bad to be cut even thick(1wm)sections.
} I am wondering if the problem come from the chemical of the set( always from EMS) since there are few labs has the same problem here.
} I appreciate any info from any body.
} Gina Zhang
}
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And to embed large parts of plant seeds - infiltration for a month or
so... time to take a break and leave your tissue sitting in resin!

cheers,
Rosemary

Dr Rosemary White
CSIRO Plant Industry
GPO Box 1600
Canberra, ACT 2601
Australia

T 61 2 6246 5475
F 61 2 6246 5334
E rosemary.white-at-csiro.au

On 21/12/12 4:19 AM, "DusevichV-at-umkc.edu" {DusevichV-at-umkc.edu} wrote:

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9, 34 -- -spurrs
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X-from: Richard Wuhrer {richard.wuhrer-at-uws.edu.au}

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Dear All,

The 12th Biennial Australian Microbeam Analysis Society Symposium (AMAS XII) will be held in at the
University of Technology, Sydney from the 4th to 8th February 2013.

The aim of the AMAS Symposium is to provide a forum where participants can discuss and share ideas
on current advances, trends and challenges in microanalysis and imaging, with an emphasis on
practical solutions and applications. We strongly encourage you to present your research work using
scanning or transmission electron microscopy and microanalysis.

A wide range of introductory and advanced workshops in the general area of microscopy and
microanalysis will be run prior to the Symposium on the 4th and 5th February 2013.

Visit www.microscopy.org.au {http://www.microscopy.org.au} for further information on; registration,
accommodation, call for papers, student travel bursary information as well as the technical and
social program.

Could you please distribute this email to your colleagues and anyone that you think might be
interested in attending.

Best regards,

Richard Wuhrer (UWS) Co-Chair

richard.wuhrer-at-uws.edu.au {mailto:richard.wuhrer-at-uws.edu.au}

*Dr Richard Wuhrer*

Research Manager

Advanced Materials Characterisation Facility

University of Western Sydney

Office of Pro-Vice Chancellor (Research)

Academic and Research Division

Locked Bag 1797 Penrith NSW 2751 Australia

*Mobile: 0411 877 476*

Phone: +61 2 9685 9089

Fax: +61 2 9685 9915

Email:*Richard.Wuhrer-at-uws.edu.au* {mailto:Richard.Wuhrer-at-uws.edu.au}

Web page: http://www.uws.edu.au {http://www.uws.edu.au/}

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I would like to know if anyone might have a standard operating procedure or
recipes for the Total Release(TM) Method for FIB lift-outs that they could
share with me. I'm interested in both plan-view and cross sections. We
have an Omniprobe 200, but it does not have the rotation option.

-Scott

Scott D. Walck, Ph.D.
100 Matte Ln
Havre de Grace, MD 21078

S.Walck-at-comcast.net
SWalck65-at-gmail.com

(760) 685-2815 (Cell)
(443) 502-5723 (Home)

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Hello All,

I have a question for the bright minds that belong to this helpful listserver here. Recently at our lab, we did EDS analysis of a Bronze alloy sample that showed Cu both K-lines and L-lines very prominently. From one point in the sample, the spectra had K-lines that were much larger than the L's. In a point just next to the previous test, the EDS spectra showed a much bigger L-line family. Any ideas of why this would happen, or what the EDS is telling us?

Thanks again,

Andrew Ornelas

Valued Quality. Delivered.
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I have no explanation for clean freshly polished bronze specimen. Otherwise difference in peak intensities could be result of higher absorbance of L peak. May be specimen has spotty organic contamination or have well developed topography.

Yesterday I got 35 "out of office" responses. Hope to beat it today.

Merry Christmas and Happy Holidays!

Vladimir

Vladimir M. Dusevich, Ph.D.
Electron Microscope Lab Director
371 School of Dentistry
650 E. 25th Street
Kansas City, MO 64108-2784

} -----Original Message-----
} From: andrew.ornelas-at-intertek.com [mailto:andrew.ornelas-at-intertek.com]
} Sent: Friday, December 21, 2012 12:11 PM
} To: Dusevich, Vladimir
} Subject: [Microscopy] EDS K-line and L-line Height
}
}
}
}
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} Hello All,
}
} I have a question for the bright minds that belong to this helpful listserver
} here. Recently at our lab, we did EDS analysis of a Bronze alloy sample that
} showed Cu both K-lines and L-lines very prominently. From one point in the
} sample, the spectra had K-lines that were much larger than the L's. In a point
} just next to the previous test, the EDS spectra showed a much bigger L-line
} family. Any ideas of why this would happen, or what the EDS is telling us?
}
} Thanks again,
}
} Andrew Ornelas
}
} Valued Quality. Delivered.
} ------------------------------------------------------------------------------
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7, 30 -- From DusevichV-at-umkc.edu Fri Dec 21 13:16:05 2012
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Wonder what the surface of the material was like? Rough?
Using sufficiently high probe voltage, } 15 kV, one thing that could
affect the relative K/L ratios is selective attenuation of lower energy
x-rays by irregular sample conditions. This could shield lower energy
x-rays more effectively than higher ones, reducing the relative L-line
intensities.

An easy way to tell if this is a factor is to examine the spectrum
backgrounds. If the spectrum with higher L-lines appears normal ( or
exaggerated), and the background of the other is lower as the x-ray
energy lowers, it is a pretty sure bet it was shielding.

Two other situations that can produce a similar effect but to a lesser
degree would be a significant change in local tilt angle or a very large
change in the average atomic number of the analysis volume.

Woody

On 12/21/2012 10:19 AM, andrew.ornelas-at-intertek.com wrote:
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} Hello All,
}
} I have a question for the bright minds that belong to this helpful listserver here. Recently at our lab, we did EDS analysis of a Bronze alloy sample that showed Cu both K-lines and L-lines very prominently. From one point in the sample, the spectra had K-lines that were much larger than the L's. In a point just next to the previous test, the EDS spectra showed a much bigger L-line family. Any ideas of why this would happen, or what the EDS is telling us?
}
} Thanks again,
}
} Andrew Ornelas
}
} Valued Quality. Delivered.

--

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Email: bsmit112-at-jhmi.edu
Name: Barbara Smith

Organization: Johns Hopkins University

Title-Subject: [Filtered] Ross Optical Lens Tissue: Supplier?

Message: We have been looking for the Ross Optical Lens Tissue and have not been able to find a
supplier. Has anyone been able to find a source or a replacement tissue? EMS shows the image of the
Ross Tissue but sends their own brand now. All the replacements we have tried are not fiber free and
lint free. Any suggestions/advice/sources would be greatly appreciated.

Thank you,
Barbara

Barbara J. Smith
EM Specialist
JHU SOM Microscope Facility
Physiology G04
725 N. Wolfe Street
Baltimore, MD 21205

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On Dec 13, 2012, at 4:36 PM, wa5ekh-at-juno.com wrote:

} I walked into a fairly new site with an older scope, which flooded
} me with memories of questions about ...well...here we go..
}
} Let's see ..that..which we don't: (help me out OEMs and techies..)
}
} 1)What temperature ranges in each em component make a long or short
} term difference?
}
} 2) Can different types pumps transmit vibrations to lenses, DPs, etc.
}
} 3) What type of chemical characteristics effect the instrument
} lines,the pumps, the hoses(!),..?
}
} 4) Does the instrument(s) or the cooling system have flow, pressure
} and temperature monitors with feedback correction and auto-shutdown
} mechanisms? Other related controls needed?
}
} 5) I have seen several cooling pump systems fail due to clogged
} filters in the system that I was not aware of..and it didn't look
} like they were intended to be checked (even hidden in the
} system)..so they just... fail.
}
} 6,7,... How many other situations are overlooked because they are
} "just cooling systems"?!
}
} I corrected and modified an SEM cooling system(changed pump design)
} once, after the 2nd pump failure failure, to find out that
} resolution/interference fluctuations dramatically disappeared. It
} occurred to me that this is more important an issue at every
} scope(high/TEM and low/dp-SEM res.)than we might have noticed.

Dear Jeff,
I'll try to answer, but there are a few items that I'm not
knowledgeable about. 1) Temperature changes degrade the image by
causing drift, but I don't know of particular short term effects of
constant temperature within a reasonable range. On the other hand,
instruments are designed to operate within a particular temperature
range, so there may well be either long or short term effects if one
operates outside the recommended range. Of course, operating
conditions that degrade such components as the epoxy used in lens
coils, plastic hoses or fittings, etc., would have bad effects. 2)
Not only can pumps transmit vibrations, but some couplings, such as
elbows, can generate turbulence, which produces vibrations. Ideally,
one wants laminar flow without pressure pulses. 3) One of the most
significant properties of cooling water that affects these components
is pH. the cooling water should be slightly basic--pH 7.5 to 8.0--in
order not to corrode copper. Distilled water, in particular, can be
quite corrosive. Since it has no dissolved ions, there is a driving
force in the direction of an equilibrium in which there are such
ions. I have been very successful using a corrosion inhibitor in
cooling systems, but check with your manufacturer and service person
to see if that is advisable for your instrument. 4) The scopes I've
worked with do have flow sensors and automatic shutdown features.
Sometimes these can become troublesome, so they must be monitored
periodically. 5) Filters are a good idea, and, once again, require
periodic monitoring and maintenance. If the ones in your scope are
hidden, I would put in additional filters in easy-to-see locations.
Again, however, check with your service person; he or she might have
been checking them and performing appropriate maintenance. 6.7...)
Proper maintenance is essential to optimal performance of any
instrument. There are no excuses because "it's just a cooling system"
or any other component. Do not let anyone cut corners on essential
maintenance for budget, time, or any other reason.
Yours,
Bill

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Barbara,
Something has happened to the Ross Lens tissue company and it is not available anymore. Ted Pella has developed a lens tissue very much like the old Ross tissue and is also in a pop-pull out box. I use it and like it very much. Hope that helps. Not sure what the order number is but I am sure if you call them they can help you with it.

Lita Duraine
Certified Electron Microscopist
Howard Hughes Medical Institute
Molecular Genetics

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Lita Duraine
7102 Gettysburg Dr.
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281-545-2054

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Andrew,

You didn't say what type of Bronze it was or what the processing treatment
was. This will determine the microstructure of the material. Others have
commented on the topography of the sample and the influence on the relative
intensities of the L lines to K lines. I suspect that you have a two phase
bronze alloy. The point with the higher Cu L/K is probably less Sn and the
point with the lower Cu L/K has higher Sn. The Sn being a higher Z material
is absorbing the L lines. Another actor could be Al. The K line for Al is
not too far above Cu L line.

What does the backscatter image look like?

-Scott

-----Original Message-----
X-from: andrew.ornelas-at-intertek.com [mailto:andrew.ornelas-at-intertek.com]
Sent: Friday, December 21, 2012 1:21 PM
To: S.Walck-at-comcast.net

Hello All,

I have a question for the bright minds that belong to this helpful
listserver here. Recently at our lab, we did EDS analysis of a Bronze alloy
sample that showed Cu both K-lines and L-lines very prominently. From one
point in the sample, the spectra had K-lines that were much larger than the
L's. In a point just next to the previous test, the EDS spectra showed a
much bigger L-line family. Any ideas of why this would happen, or what the
EDS is telling us?

Thanks again,

Andrew Ornelas

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As others have suggested, etching stainless steel can be
tricky. Surface preparation is critical, especially for
small wire with cold worked structures. The Vandervoort book
is a great reference with many options. The ASM Handbooks on
Metallography (Vol 9, I think) are also excellent, and one
or more editions of this set will likely be in your library.
One issue that you might have with these samples, since you
are a neophyte in this area, is recognizing when you have
revealed the true structure. A small wire may have such
highly elongated grains that all you may see with a perfect
preparation and etch is a jumble of lines generally parallel
to the wire.

Anyway, back to the etchant question. For type 316 stainless
steel, picric acid solutions are not going to work nor are
Kalling's or Marble's reagents likely to be very successful.
Even the classic electrolytic etch with 10% oxalic acid for
austentic stainless alloys can be less than satisfactory for
T316. Here are some options for you:

1. If you want general grain structure, electrolytic etching
at about 6 VDC in 10% ammonium persulfate in water will
usually give a more uniform and reliable etch than oxalic
acid for T316.

2. An immersion etch for general structure (if you don't or
can't do electrolytic) is a solution of 12.5 g of CuCl2 in
350 ml of concentrated hydrochloric acid mixed at a 3:1
ratio with concentrated nitric acid (the CuCl2 and HCl can
be mixed and stored, but only mix in the nitric just before
etching). Immerse or swab gently for a few seconds. This
etch is sensitive to surface conditions; a good polish is
essential and etch immediately after polishing. (The
stainless steel polished surface will passivate quickly in
ambient air, which inhibits etching.)

3. If you want only to see the grain boundaries, try
electrolytic etching with 60% nitric acid in water at 1.1
VDC (30 to 120 s).

Good luck.

On 12/18/2012 3:49 PM, edelmare-at-miamioh.edu wrote:
} To start - I'm a neophyte in metalographic crystalography.
}
} O.k., I have a user we're looking for grain structure size changes in stainless
} steel 316. Samples are some wires embedded in phenoloic resin for
} polishing. Their polished pretty well. Following a recommmendation we've
} etched with a solution of 4g picric acid in 100ml ethanol. But we are not
} seeing the grain structure in reflected light microscopy. (Moving to SEM later
} today )
}
} Is this a good etchant for SS 314? How long would you recommend etching
} for?
}
} Any other suggestions or recommendations?
}
} Thank you one and all!
}
}
} Richard E. Edelmann, Ph.D., Director
} Center for Advanced Microscopy & Imaging
} 9C Upham Hall
} Miami University, Oxford, OH 45056
} Ph: 513.529.5712 Fax: 513.529.4243
} E-mail:edelmare-at-muohio.edu
} http://www.cami.muohio.edu

--
________
Larry D. Hanke, P.E.
Principal Engineer
Materials Evaluation and Engineering, Inc.
(763) 449-8870

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Dear all,

I would like to draw your attention to a newly announced EM facility manager position in Paul Scherrer Institute, Switzerland. Please refer to
http://www.psi.ch/pa/offenestellen/0487-1

Best regards,
Takashi Ishikawa

Takashi Ishikawa
Senior Scientist
BMR, BIO
Paul Scherrer Institute (PSI)
OFLB/010
5232 Villigen PSI, Switzerland
Tel: +41 (0) 56 310 4217
Fax: +41 (0) 56 310 5288
e-mail: takashi.ishikawa-at-psi.ch
web: http://www.psi.ch/lbr/takashi-ishikawa

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Email: michael.cammer-at-med.nyu.edu
Name: Michael Cammer

Organization: NYU Langone Medical Center

Title-Subject: [Filtered] new TIRF microscopy evaluations

Message: We are looking at new TIRF microscope systems and are seeking info from users.

We are familiar with the first generations of Olympus's TIRF arm and Nikon's current implementation
on the EclipseTi with NIS Elements software and are now looking at a potential purchase of a new system.

Of particular interest, we are seeking reports/opinions about the Olympus CELL^TIRF system operating
with Olympus's software and if you have used the IX83 with the newest generation autofocus system,
info on this would be greatly helpful. Comparison with Nikon of Zeiss's current products would be
even better.

Probably personal email or off-line discussion would be preferable.

Thank you!!

________________________________________________________
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Email: a.chuvilin-at-nanogune.eu
Name: Andrey Chuvilin

Organization: CIC nanoGUNE, Ikerbasque

Title-Subject: [Filtered] FIB: Postdoc position opened in San Sebastian

Message: Dear Listers,
This is to announce an immediate opening of a postdoc position at CIC nanoGUNE, San Sebastian, Spain.

Details of the position can be found at our website:
http://www.nanogune.eu/en/research/electron-microscopy/available-positions/post-doc-position-focused-ion-beam-nanofabrication-and-related-techniques/

I will be glad to answer any additional questions concerning this offer, however applications should
be send to the address specified on the web page.

All the best and Happy New Year!

Andrey

_____________________
Andrey Chuvilin
Ikerbasque Research Professor
Electron Microscopy Laboratory
CIC nanoGUNE Consolider
Tolosa Hiribidea, 76
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Dear interested Colleague,
I'd like to inform you today of NEW CONTENT on the SCUR-Website, regarding our next Meeting in Salzburg, Austria,
hoping you'll find all information needed or you are expecting to find.

==============================================================
Congress Announcement (2nd announcement, Call for papers)
==================================================================================
40th Annual Meeting of the Society for Cutaneous Ultrastructure Research (SCUR) and
6th Joint Meeting of the Society for Skin Structure Research (SSSR, Japan)

(Affiliated Meeting with /after IID-2013 Edinburgh, UK)

"From Genetics to Therapy"

SALZBURG (AUSTRIA), May 12-14, 2013

visit: http://orgs.dermis.net/scur/scur2013/content/e01home/e01home/index_ger.html

==================================================================================

If there are question left, please let me have your request by e-mail to w.muss-at-salk.at or scursssr.salzbg2013-at-Gmail.com

Apologize if this or similar message is received in your mail-box a second time.

Happy Holidays everywhere and with your families,
wishing you and your loved ones a season filled with wonder and simple joys,
looking forward to sharing an exciting New Year,
as well as (eventually) your participation in our Meeting in May 2013!

Warmest regards
yours respectfully, but cordially too,

Wolfgang

======================
Wolfgang MUSS, PhD
Secretary of SCUR
Member of MSA
SALZBURG - AUSTRIA
www.scur.org [Meetings - Next Meeting - URL "40th Annual Meeting of SCUR = 6th Joint Meeting with SSSR 2013: SALZBURG, Austria (May 12-14, 2013")

The Website will be updated again after the holidays.

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