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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 1 Jan 2013 10:45:00 -0600
Subject: [Microscopy] Administrivia: Happy New Year 2013

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Happy New Year Colleagues;

Welcome to the 21st year of operation of the Microscopy ListServer
a free service to the world wide microscopy community, sponsored
jointly by your Friendly Neighborhood SysOp and the Microscopy Society
of America.

During 2012, the ListServer delivered 1350 messages to nearly 3500 subscribers
around the world, with minimal hassels (that I know about). For those of you that are
statistics junkies this year you generated 245+ Gbits of Email traffic and over
4.7 Million Email messages were sent out this year by my tired little server.

As usual you don't want to know how much Junk Mail and spam has been filtered out.


The complete Microscopy ListServer Archives for 2012-1993 (~ are on-line at

http://www.microscopy.com.

A couple of IMPORTANT reminders:

If you leave on vacation/holiday use the on-line form to UNSUBSCRIBE
your Email address from the listserver. The out-of-office / on-vacation
autoreply messages are a real nuisance to posters.

Do not reply to message with the return address of:

MicroscopyListserver-noreply-at-microscopy.com

these are messages forwarded usually from the WWW posting form. They do not
go back to the poster but rather into a black hole, which I rarely check.
If you see a message that has this "No-Reply" return address please post your
reply/comment/answer to:

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or if you wish to reply privately, look at the username in the body of
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As always if you have questions about suitability of postings or
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Cheers,

Nestor
Your Friendly Neighborhood SysOp


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From: schooley-at-mcn.org
Date: Tue, 1 Jan 2013 11:39:36 -0600
Subject: [Microscopy] Re: turning 21

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

} ----------------------------------------------------------------------------
} The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

Thank YOU, Ness, for all that you've done for us. Now that you're
21, you can have a legal beer to celebrate...

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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3, 18 -- From: Caroline Schooley {schooley-at-mcn.org}
3, 18 -- Subject: Re: [Microscopy] turning 21
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From: vcrvince-at-comcast.net
Date: Tue, 1 Jan 2013 15:30:03 -0600
Subject: [Microscopy] Re: turning 21

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hear, hear and I may add that Nestor's list server preceded Facebook and
LinkedIn

Vincent Carlino

vince.carlino-at-ibssgroup.com
www.ibssgroup.com



-----Original Message-----
X-from: schooley-at-mcn.org [mailto:schooley-at-mcn.org]
Sent: Tuesday, January 01, 2013 9:44 AM
To: vcrvince-at-comcast.net

} -----------------------------------------------------------------------
} ----- The Microscopy ListServer -- CoSponsor: The Microscopy Society
} of America To Subscribe/Unsubscribe --
} http://www.microscopy.com/MicroscopyListserver

Thank YOU, Ness, for all that you've done for us. Now that you're 21, you
can have a legal beer to celebrate...

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
===


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From: tina-at-pbrc.hawaii.edu
Date: Tue, 1 Jan 2013 20:27:58 -0600
Subject: [Microscopy] Re: Administrivia: Happy New Year 2013

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Ness, let me echo Caroline's sentiments! And have a beer on it (I'm making
a new batch of beer today).

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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From: oshel1pe-at-cmich.edu
Date: Wed, 2 Jan 2013 10:03:47 -0600
Subject: [Microscopy] Ask-A-Microscopist HRTEM image processing & FFT

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

***************************************************************************************
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 - Ken Hayes
} Email - khayes-at-firstsolar.com
} EDUCATION - Graduate College
} SUBJECT_OF_QUESTION - HRTEM Image Processing
} QUESTION - I collect a large number of HRTEM images in my job. I
} currently use ImageJ to view and process them. I am looking for a
} user friendly package softare package for HRTEM image processing
} with an emphesis on FFT pattern solving. the OS is Windows 7. Can
} you direct me to a list of recommended softare?
} Thanks.


--

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From: spurgeon-at-drexel.edu
Date: Wed, 2 Jan 2013 11:05:40 -0600
Subject: [Microscopy] Cross Correlation Software for STEM-HAADF Images

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi everyone,

I have a series of STEM-HAADF images taken at 40 micro-sec time
intervals that I would like to cross-correlate and average. Is there
any freely available software to do this, such as a plugin for ImageJ
or DigitalMicrograph? Also, does anyone know of a script to batch
export a series of slices from an image in DM?

I haven't had much like finding anything and would great appreciate
your suggestions.

Thanks!
--
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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From: ben.micklem-at-pharm.ox.ac.uk
Date: Thu, 3 Jan 2013 04:42:05 -0600
Subject: [Microscopy] Nikon 1 series cameras on microscopes

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear List,

I am very interested in hearing from anyone who has mounted a Nikon 1
series compact system camera on a light microscope.

I am considering trying a Nikon V1 camera and a cheap LCD monitor via HDMI
for a low-budget imaging system on an old Leitz 160mm tube length
microscope. I have a trinocular head (38mm photoport), and an adapter for
it to C-mount (with no optics in it). Is this adapter the 1x, which I
understand is suitable for covering a 1" sensor?

If so, all I need is a c-mount to 1 series mount adapter, and I should be
ready to go?

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/}





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12, 30 -- Subject: Nikon 1 series cameras on microscopes
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From: kraftpiano-at-gmail.com
Date: Fri, 4 Jan 2013 09:55:31 -0600
Subject: [Microscopy] TEM: Manual request.

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Greetings and happy new year to everyone! (And congrats, Nestor, on
being legal- have an extra one on me!)

I was wondering if anyone had a manual for a JEOL JEM-1400 TEM that I
could get a copy of? I'd be happy to pay round-trip overnight
shipping or copy costs, whichever is easier...

Thank you,

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

==============================Original Headers==============================
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From: mmcgough-at-histochemicalsociety.org
Date: Fri, 4 Jan 2013 13:05:41 -0600
Subject: [Microscopy] HCS Immunohistochemistry and Microscopy Course - Second Announcement

Contents Retrieved from Microscopy Listserver Archives
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Hello All,

There is still one week left to register for The Histochemical Society
course in Immunohistochemistry and Microscopy at MBL in Woods Hole, MA.
The deadline is JANUARY 11, 2013.

Application and complete information on website:
http://hermes.mbl.edu/education/courses/special_topics/ihcm.html

The course is four full days and evenings (11 hours daily) of lecture
and laboratory sessions with experts in the field of
immunohistochemistry (IHC) and microscopy.

FASEB/MARC AND HCS TRAVEL AWARDS for THE COURSE

The FASEB MARC Program provides funding for travel awards to encourage
and support the participation of underrepresented students and
postdoctoral fellows in the HCS IHCM course. The deadline for applying
is February 9, 2013:
http://www.faseb.org/MARC-and-Professional-Development/Travel-Awards.aspx

The Histochemical Society offers travel awards to attend the HCS Course.
The deadline for applying is January 31, 2013:
http://histochemicalsociety.org/Awards/Students---Post-Docs.aspx

This is a fabulous opportunity, click on the links and apply now!

Kind regards,
Meg McGough
mmcgough-at-histochemicalsociety.org


==============================Original Headers==============================
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From: sergei2-at-ornl.gov
Date: Fri, 4 Jan 2013 13:31:09 -0600
Subject: [Microscopy] PFM School at ORNL - March 4-8, 2013

Contents Retrieved from Microscopy Listserver Archives
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Dear colleagues - I would like to bring to your attention a *12th
International Workshop on Piezoresponse Force Microscopy and
Electrochemical Strain Microscopy, *to be held at The Center for
Nanophase Materials Sciences, Oak Ridge National Laboratory, on March
4-8 this year. This workshop is organized by S.V Kalinin, A.P. Baddorf,
and A. Kholkin, and will feature a set of tutorials on advanced PFM and
ESM operation, and extensive hands-on labs. The attendance will be
limited to ~24 people. Please see below the information on the scope and
content of the school.

Yours truly

Sergei V. Kalinin

Piezoresponse Force Microscopy (PFM) has by now become one of the
dominant techniques for exploring ferroelectric, multiferroic, polar
macromolecular and biological, and ionic materials on the nanoscale. The
last several years have seen multiple breakthroughs in practice and
theory of PFM achieved in multiple research groups worldwide. This
workshop aims to provide an in-depth description of recent advances in
Piezoresponse Force Microscopy and to offer laboratory demonstrations
designed for advanced PFM practitioners. The workshop will introduce
basic principles of PFM operation, relevant instrumental aspects, and
image interpretation. The theory of cantilever dynamics, PFM contact
mechanics, and resolution theory, as well as their implications for
qualitative and quantitative data interpretation in PFM, will be
presented and illustrated experimentally. Recent technical advances in
PFM, including vector PFM, high-frequency PFM, band-excitation and dual
frequency resonancy tracking (DFRT) imaging, switching spectroscopy PFM
and imaging and polarization switching in liquids and vacuum, will be
presented. For ferroelectric materials, applications of PFM for domain
imaging, nucleation center mapping, and probing polarization dynamics in
thin films and capacitor structures will be discussed and demonstrated.
Finally, electromechanical probing of biological, electroactive polymer,
and soft-condensed matter systems beyond classical ferroelectric
applications will be described.

The 4 day workshop designed for advanced PFM users will include
tutorials and lectures given by leading PFM experts and experimental
hands-on tutorials on PFM imaging and spectroscopy. Ultimately, the goal
of the workshop is to build a network of advanced PFM practitioners to
promote rapid dissemination of theoretical knowledge, experimental
protocols, and novel technique development in this rapidly growing area,
as well as to establish links to areas such as energy storage and
conversion and information technology, macromolecular science and
electrochemistry.

The workshop is supported by the Center for Nanophase Materials Sciences
at Oak Ridge National Laboratory and the European Initial Training
Network (ITN) ”Nanomotion.” Contact Sergei V. Kalinin (sergei2-at-ornl.gov
{mailto:sergei2-at-ornl.gov} ) for details.


**

*Piezoresponse Force Microscopy: Theory, Techniques, and Applications*

The workshop will open with a series of the tutorial talks on principles
and applications of PFM in the mornings and series of hands-on
experiments (samples brought by attendees) in the afternoon. The
hands-on training will be performed by the CNMS staff members of extant
platforms for the groups of ~3 people.

*Tentative list of tutorials:*

1. (Kholkin) Basic principles and history of PFM
2. (Kalinin) Contact mechanics, resolution theory, and interpretation of
PFM signal
3. (Kalinin) Cantilever dynamics and frequency dependent measurements in PFM
4. (Kalinin) Polarization switching in PFM: voltage and time spectroscopies
5. (Jesse) Band excitation methods and BE PFM
6. (Belianinov) Multivariate statistical methods: working with
multidimensional data
7. (Kalinin) Advanced PFM modes: spectroscopy and multivariate analysis
8. (Rodriguez) PFM and KPFM in liquids
9. (Maksymovych) Combined PFM and cAFM: ferroelectric tunneling and
domain walls
10. (Balke) Electrochemical Strain microscopy of Li-ion materials
11. (Kalinin) ESM of oxygen conductors and electrochemical effects in PFM
12. (Shvartsman) PFM of relaxor ferroelectrics
13. (Rodriguez) PFM of ferroelectric capacitors
14. (Martin) Domain structures and polarization dynamics in PZT and BFO
15. (Rodriguez) PFM of macromolecular and biological systems
16. (Jesse) Local thermal characterization
17. (Tselev) Microwave imaging


==============================Original Headers==============================
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From: FMonson-at-wcupa.edu
Date: Sun, 6 Jan 2013 01:44:56 -0600
Subject: [Microscopy] Ask-A-Microscopist HRTEM image processing & FFT

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Might try the following.

http://reindeergraphics.com/

Chrs&HNY,

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
________________________________________
X-from: oshel1pe-at-cmich.edu [oshel1pe-at-cmich.edu]
Sent: Wednesday, January 02, 2013 11:12 AM
To: Monson, Frederick

***************************************************************************************
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 - Ken Hayes
} Email - khayes-at-firstsolar.com
} EDUCATION - Graduate College
} SUBJECT_OF_QUESTION - HRTEM Image Processing
} QUESTION - I collect a large number of HRTEM images in my job. I
} currently use ImageJ to view and process them. I am looking for a
} user friendly package softare package for HRTEM image processing
} with an emphesis on FFT pattern solving. the OS is Windows 7. Can
} you direct me to a list of recommended softare?
} Thanks.


--

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From: petereaton-at-hotmail.com
Date: Tue, 8 Jan 2013 04:55:48 -0600
Subject: [Microscopy] AFM short training course

Contents Retrieved from Microscopy Listserver Archives
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Dear Colleagues,
We are running a four day training course in Atomic Force Microscopy over Easter this year.
This is the second time we've run the course, and the students were .very positive about the first edition.
The course will be held in our laboratory in Porto, Portugal, with two instruments for the students to use. The course will include days of theory and two days of practice, covering both Image acquisition and data analysis. The course will take place between the 25th and the 28th of March 2013.2. Places are very  limited with just a few remaining, so interested students are encouraged to reserve a place as soon as possible, but definitely before 5th January. 
Visit http://bit.ly/TuuO2 for more information. Enquiries and reservations can be made by emailing afmhelp-at-gmail.com

____________________________________________________________________________
Atomic Force Microscopy
Dr Peter Eaton and Dr Paul West
Available through all good bookshops, or direct from Oxford University Press at:
http://ukcatalogue.oup.com/product/780199570454..do http://afmhelp.com



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4, 19 -- Subject: AFM short training course
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From: jerrysedgewick-at-gmail.com
Date: Wed, 9 Jan 2013 09:03:42 -0600
Subject: [Microscopy] Re: Nikon 1 series cameras on microscopes

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html


Dear Colleagues, Apologies for the previous email which said that application for our AFM training course must be made by January the 5th. There was were some formatting errors in the email. The correct deadline is *January 15th*.
Regards,
Pete.



____________________________________________________________________________
Atomic Force Microscopy
Dr Peter Eaton and Dr Paul West
Available through all good bookshops, or direct from Oxford University Press at:
http://ukcatalogue.oup.com/product/9780199570454.do http://afmhelp.com



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Message-ID: {C47C213BBBB5A44F8E6D98AED28167F893756A48-at-speed-magazin.com}

Hi Ben,

I've done this with a Canon camera. These are a bit easier because the
phototube can be purchased, and Canon has built in features for
tethering the camera to the computer, live focus, etc.

However, it looks like you're on the right track. The sensor has to be
160mm from the back of the objectives. Otherwise, the focus point will
change for the objective when a specimen is focused by eye. It's best to
devise a means to slide the camera up and down along 2 tubes (a wider
diameter tube and a narrower diameter tube), or to order a focusing tube
from Thorlabs. The quick and dirty way is to use one tube at a narrower
diameter and a larger diameter "sleeve." Have a machine shop drill
threaded holes into the sleeve for thumb screws (or, if you're like me,
you do it yourself with a drill press and hand threading device). It may
be a bit difficult to get two different sized tubes that are meant to
fit to each other, but you can try Thorlabs or Edmund Optics.

Mount your camera, focus by eye, and then, while the camera is giving
you a live view, adjust the outer sleeve up or down until the image is
in focus. Then tighten the thumb screws against the inner tube at that
position.

The image may not fill the sensor. Who cares. I suspect you will have
plenty of pixel coverage, even for Nyquist rates. Simply crop the images
after these are acquired, best done via an automated routine in
Photoshop or other imaging program. In that way, the crop will always
remain the same. Anyway, you'll likely get darkening at the edges
(vignetting), some barrel distortion, out-of-focus areas, etc., so even
if the entire sensor is filled, you'll still want to exclude image edges.

If it goes the other way and the sensor is overfilled, then you will
have to drop a lens into the tube.

If you could find out the focal length of the tube lens in that model of
Leica, then you could calculate coverage of the sensor (there's a lens
after the objective along the light path). Or, you can turn off all the
lights and hold a piece of paper where the sensor *should* be to get the
diameter of the virtual image when focused.

Good luck!

Jerry




On 1/3/2013 4:50 AM, ben.micklem-at-pharm.ox.ac.uk wrote:
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} Dear List,
}
} I am very interested in hearing from anyone who has mounted a Nikon 1
} series compact system camera on a light microscope.
}
} I am considering trying a Nikon V1 camera and a cheap LCD monitor via HDMI
} for a low-budget imaging system on an old Leitz 160mm tube length
} microscope. I have a trinocular head (38mm photoport), and an adapter for
} it to C-mount (with no optics in it). Is this adapter the 1x, which I
} understand is suitable for covering a 1" sensor?
}
} If so, all I need is a c-mount to 1 series mount adapter, and I should be
} ready to go?
}
} 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/}
}
}
}
}
}
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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
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From: oshel1pe-at-cmich.edu
Date: Wed, 9 Jan 2013 12:53:28 -0600
Subject: [Microscopy] Ask-A-Microscopist Lowicryl sectioning problems

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***************************************************************************************
Forwarded from "Ask a Microscopist"
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} realname - Ekaterina
} Email - Eyarunov-at-uwyo.edu
} EDUCATION - Graduate College
} QUESTION - Hello! I am having problems with sectioning my blocks for
} TEM. I have blocks of cells embedded into Lowicryl resin and I am
} trying to obtain sections 50-70 nm thick. However my sections are
} very wrinkled and I cannot use them for further immunoEM. What would
} be the reasons that can be fixed to avoid wrinkling? Temperature,
} speed of cutting, etc?
} Thank you for you help,
} Ekaterina


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From: mmcgough-at-histochemicalsociety.org
Date: Thu, 10 Jan 2013 16:09:41 -0600
Subject: [Microscopy] HCS Immunohistochemistry and Microscopy Course Registration Extended

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Hello All,

Registration for The Histochemical Society course in
Immunohistochemistry and Microscopy at MBL in Woods Hole, MA has been
extended until JANUARY 25, 2013.

Application and complete information on website:
http://hermes.mbl.edu/education/courses/special_topics/ihcm.html

The course is four full days and evenings (11 hours daily) of lecture
and laboratory sessions with experts in the field of
immunohistochemistry (IHC) and microscopy.

FASEB/MARC AND HCS TRAVEL AWARDS for THE COURSE

The FASEB MARC Program provides funding for travel awards to encourage
and support the participation of underrepresented students and
postdoctoral fellows in the HCS IHCM course. The deadline for applying
is February 9, 2013:
http://www.faseb.org/MARC-and-Professional-Development/Travel-Awards.aspx

The Histochemical Society offers travel awards to attend the HCS Course.
The deadline for applying is January 31, 2013:
http://histochemicalsociety.org/Awards/Students---Post-Docs.aspx

Please email if there are any questions, happy to help.

Kind regards,
Meg McGough
mmcgough-at-histochemicalsociety.org

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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 12 Jan 2013 07:53:10 -0600
Subject: [Microscopy] viaWWW:Averaging STEM-HAADF images

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Email: LES-at-ZSGENETICS.COM
Name: Larry Scipioni

Organization: ZS Genetics

Title-Subject: [Filtered] Averaging STEM-HAADF images

Message: Dear Steven,
If you are looking to align and average a stack of noisy images, StackReg works great in ImageJ.
Then just use the "Z Project" command to average it all.
Also, if you use "Save as" tif in DM on a stack, it will produce a multipage tif that opens up
directly in ImageJ as a stack, ready for averaging.


Steven Spurgeon wrote:

I have a series of STEM-HAADF images taken at 40 micro-sec time
intervals that I would like to cross-correlate and average. Is there
any freely available software to do this, such as a plugin for ImageJ
or DigitalMicrograph? Also, does anyone know of a script to batch
export a series of slices from an image in DM?


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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 12 Jan 2013 07:55:22 -0600
Subject: [Microscopy] viaWWW:Leitz 1512 Microtome

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Email: tkremer-at-ipstesting.com
Name: Tom Kremer

Organization: IPS Testing

Title-Subject: [Filtered] Leitz 1512 Microtome

Message: I inherited a Leitz 1512 rotary microtome. It's in excellent shape but without any knives.
On occasion I will prepare cross sections of embedded polymer fiber networks, polymer films and wood
fiber based structures. Would some of you who have used or maybe still use this tool give me some
recommendations on knives, where you prefer to have them sharpened and preferred knife orientation
for optimum cutting? I would appreciate it.

Tom Kremer
IPS Testing

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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 12 Jan 2013 07:56:35 -0600
Subject: [Microscopy] viaWWW:need help identifying unknown bug

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Name: David Lowry

Organization: Arizona State Univ

Title-Subject: [Filtered] need help identifying unknown bug

Message: we are seeking help from list-members with microbiology background in identifying an
endoparasite that is causing contamination problems in cultures of green algae. The contaminant
appears to be a bacterium but possesses an unusual morphology and apparent mode of reproduction. If
you might be able to assist, please contact me off-line and I can forward a few representative
images for evaluation. thank you-

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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 12 Jan 2013 07:57:23 -0600
Subject: [Microscopy] viaWWW:Ultracut S control box shut down

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Name: Mary Ard

Organization: University of Georgia, College of Veterinary Medicine

Title-Subject: [Filtered] Ultracut S control box shut down

Message: I have a Reichert Ultracut S ultramicrotome which has served me well for many years with
little to no problems until yesterday. The control box shut down completely. I have checked all
outside connections and have replaced the fuse (which I found blown - 220v). I'm sure it is an
internal problem, but I hesitate to open the box without an expert opinion. When replacing with
another 220v fuse, I can feel some activity coming from the inside, but the control box does not
come on. When replacing with a 110v fuse, the box hums, the initial light display comes on, but
then shuts down. Has anyone had a similar experience with this?

Regards,
Mary

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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 12 Jan 2013 07:58:36 -0600
Subject: [Microscopy] viaWWW:We are buying a Dual Beam FIB

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Email: jefbettini-at-gmail.com
Name: Jefferson Bettini

Organization: Brazilian Nanotechnology National Laboratory (LNNano)

Title-Subject: [Filtered] We are buying a Dual Beam FIB

Message: This laboratory is currently evaluating the purchase of a dual beam FIB and I recently
learned that Carlos Kazuo Inoki, who is an employee from this laboratory, posted some comments about
products from FEI, ZEISS and other companies, in internet sites. These comments represent
exclusively the personal opinion from Carlos Inoki and they do not express any institutional
assessment or opinion of the Brazilian Nanotechnology National Laboratory (LNNano), represented by
its management. Indeed, members of this laboratory have a completely different opinion from Carlos
Inoki.
LNNano management and personnel have been in contact with different companies concerning this
acquisition and we welcome their proposals since this will allow us to reach a sound decision,
considering the needs of our users and researchers. Finally, we acknowledge JEOL and FEI for all the
support these companies have provided to their microscopes currently installed in this laboratory.



Jefferson Bettini
Facillity Manager, Electron Microscopy Laboratory
Brazilian Nanotechnology National Laboratory (LNNano)


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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 12 Jan 2013 07:59:42 -0600
Subject: [Microscopy] viaWWW:water-solvent swap

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Name: Marissa

Organization: LBNL

Title-Subject: [Filtered] water-solvent swap

Message: I have nanoparticles in what I suspect to be dirty water and would like to transfer them
into a clean solvent for drop-casting. Is it possible to evaporate the water and then add the
appropriate solvent or am I completely oblivious to some well known nanoparticle in solution rule?

Thanks!

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From: dsherman-at-purdue.edu
Date: Sat, 12 Jan 2013 08:32:16 -0600
Subject: [Microscopy] Re: viaWWW:water-solvent swap

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Marissa,

Evaporating the water will only concentrate the impurities and will likely
aggregate the nano particles as well. You need to look at methods using
filtration or centrifugation. However, I would start by looking at the
method used to prepare the nano particles initially to see if there are
ways to prevent the contamination in the first place.

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
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From: jpapalia-at-papalia.net
Date: Sat, 12 Jan 2013 18:52:24 -0600
Subject: [Microscopy] Re: viaWWW:water-solvent swap

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Hi Marissa,

There's a lot to consider when contemplating cleaning up nanoparticles
in solution. Do the nanoparticles have any form of functionalization?
A peptide group? A surfactant? Anything? or do they just have an
appropriate counterion in the solution which keeps them from
flocculating/agglomerating? The answer there will determine your next
step, and whether or no you can easily consider changing solvents.

Assuming your particles have something (surfactant or otherwise) to
prevent agglomeration, then you can consider a centrifugation / wash /
re-suspend / (sonication?) / centrifugation... routine.

Depending on the weight/size of your nanoparticles, you can also
consider using dialysis tubing, as long as there is a significant enough
weight difference between your impurity and your nanoparticles. That
will also determine the diffusing species and direction.

Once you get past those questions, then you can start to consider
whether or not you can even suspend the nanoparticles in your desired
solvent.

-John

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}
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} into a clean solvent for drop-casting. Is it possible to evaporate the water and then add the
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From: microscopylistserver-noreply-at-microscopy.com
Date: Sun, 13 Jan 2013 09:31:43 -0600
Subject: [Microscopy] viaWWW:Post-Doctoral Research Associate in Focussed Ion Beam (FIB)/Electron

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Name: Richard Harrison

Organization: University of Cambridge

Title-Subject: [Filtered] Post-Doctoral Research Associate in Focussed Ion Beam (FIB)/Electron
Microscopy, University of Cambridge

Message: Post-Doctoral Research Associate in Focussed Ion Beam (FIB)/Electron Microscopy

Applications are invited for a post-doctoral research associate (PDRA) position within the mineral
magnetism group of the Department of Earth Sciences, University of Cambridge. The position is funded
by the ERC grant “Nanopaleomagnetism: a new multiscale approach to paleomagnetic analysis of
geological materials”, a brief description of which is given below. Funding is available for 3 years
in the first instance. Successful applicants will hold a PhD on taking up the post.

The PDRA project will focus on the application of a dual-beam focussed ion beam (FIB) workstation to
perform 3D slice-and-view tomography of natural samples. The person appointed will use facilities
located in the Department of Materials Science to study the positions, sizes, shapes, compositions
and crystallographic orientations of magnetic particles embedded in silicate hosts using a
combination of slice-and-view tomography, electron backscattered diffraction (EBSD) and energy
dispersive X-ray analysis (EDX). We seek candidates with a strong background in electron microscopy
and specific experience with using FIB techniques. Experience with EDX, EBSD and tomography would be
an advantage. It is not necessary to have a background in either Earth sciences or magnetism, and we
invite suitably experienced candidates from any area of the physical sciences. The successful
candidate will have a track record of publication in peer-reviewed journals, will have good
communication skills and have demonstrated the capacity to perform independent research.

For full details see http://www.jobs.cam.ac.uk/job/-24782/

Interested candidates should contact Dr. Richard Harrison (rjh40-at-esc.cam.ac.uk) for more information
about the project and formal application procedures.

General description of the ERC project:

Adopting cutting-edge techniques from physics and materials science, Nanopaleomagnetism aims to
perform paleomagnetic measurements at submicron length scales, enabling primary magnetic signals to
be extracted from ancient and severely altered geological materials. 3D measurements of the volume,
shape and spacing of all magnetic particles within a microscale region of interest will be made
using a ‘dual beam’ focussed ion beam workstation. Combined with high-resolution paleomagnetic
measurements and nanometre/nanosecond electron/X-ray magnetic imaging, nanopaleomagnetism will, for
the very first time, be able to characterise the magnetic properties of geological materials at
fundamental length scales and time scales. The nanoscale measurements will enable us to capture the
essential physics of the remanence acquisition process and to explore magnetic behaviour ‘in
silicoÂ’, allowing predictions to be made that can be tested directly against experimental
observations at all length scales. Sample-return missions to asteroids, comets, moons and planets
will soon provide unprecedented opportunities for extraterrestrial paleomagnetism.
Nanopaleomagnetism will provide the methodology and instrumentation needed to analyse these precious
materials.

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From: hyi-at-emory.edu
Date: Sun, 13 Jan 2013 11:45:35 -0600
Subject: [Microscopy] Lowicryl polymerization

Contents Retrieved from Microscopy Listserver Archives
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Dear Researchers:

I am doing a cryosubstitution run. The samples being processed need to
be positioned in a certain way for Lowicryl embedding. The process of
positioning samples takes time and needs to be done under a dissecting
scope, therefore it would be easier if it is done when the samples are at
a higher temperature (near zero), and subsequently embed/UV polymerize in
Lowicryl at the same temperature. Has anyone tried this? Is there
anything I need to be aware when UV polymerize Lowicryl at this
temperature? Thanks in advance.

Hong



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9, 38 -- Subject: Lowicryl polymerization
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From: hyi-at-emory.edu
Date: Sun, 13 Jan 2013 11:50:55 -0600
Subject: [Microscopy] JEOL JEM-1400 TEM

Contents Retrieved from Microscopy Listserver Archives
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Dear Researchers:

I would like to get some information about JEOL JEM-1400 TEM. If you own
this microscope, I would be delighted to hear from you off-line. Thank
you very much in advance.

Hong


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From: dsherman-at-purdue.edu
Date: Sun, 13 Jan 2013 14:11:32 -0600
Subject: [Microscopy] Re: Lowicryl polymerization

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hong,

I have often polymerized Lowacryl HM20 using UV light at 0oC or -20oC. In
fact, for years we used a makeshift UV setup in a refrigerator freezer. I
just cut out poster board and covered them with aluminum foil to make a
reflective chamber. I had two plexiglass rods that were poked through the
side pieces and supported a test tube rack. The bottom wires on the rack
were just the right size for suspending Beem capsules by their lids. It
also would hold flat molds if necessary. All the actual orientation was
done in the flat mold on a cold tray using a dissecting microscope.In that
case the mold on the cold tray could be moved to the reflective chamber
and UV light suspended from above.

Later we did get a Leica FS unit with the binocular attachment. This
meant that we could orient in flat molds while still at very low
temperatures and then not have to move the mold until after
polymerization. That works great so we could then polymerize at much
lower temperatures. I found that many antigens were preserved just fine
at 0 or -20 but then there are those elusive few where labeling really
seemed to benefit from lower temperatures.

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
www.dsimagingllc.com
Mobile: 765-418-8540



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From: microscopylistserver-noreply-at-microscopy.com
Date: Sun, 13 Jan 2013 23:53:36 -0600
Subject: [Microscopy] viaWWW:Porter Blum MT2 Ultramicrotome Manual/Parts?

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Email: wa5ekh-at-juno.com
Name: Jeff Day

Organization: Texas Industrial Investments/ JD- Sole Investor/Microscopy Advisor

Title-Subject: [Filtered] Porter Blum MT2 Ultramicrotome Manual/Parts?

Message: I bought a MT2, and, of course, I need parts and a manual(hard or electronic). Can anyone
suggest some sources?

Anyone using one these days? How do they still run?

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From: ehaller-at-health.usf.edu
Date: Mon, 14 Jan 2013 07:25:27 -0600
Subject: [Microscopy] Lowicryl polymerization

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi, Hong,

Is it possible to do a partial orientation on the samples, that is, to orient them in one direction, and polymerize them in the Lowicryl at the usual temperature, then cut the polymerized blocks with a saw, and Super-Glue the cut blocks onto other blocks to obtain your final orientation for sectioning?

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: hyi-at-emory.edu [hyi-at-emory.edu]
Sent: Sunday, January 13, 2013 12:52 PM
To: Haller, Edward

Dear Researchers:

I am doing a cryosubstitution run. The samples being processed need to
be positioned in a certain way for Lowicryl embedding. The process of
positioning samples takes time and needs to be done under a dissecting
scope, therefore it would be easier if it is done when the samples are at
a higher temperature (near zero), and subsequently embed/UV polymerize in
Lowicryl at the same temperature. Has anyone tried this? Is there
anything I need to be aware when UV polymerize Lowicryl at this
temperature? Thanks in advance.

Hong



________________________________

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From: PWebster-at-hei.org
Date: Mon, 14 Jan 2013 09:32:12 -0600
Subject: [Microscopy] Lowicryl polymerization

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html


Dear All,

With Lowicryl, it is also possible to embed using UV illumination at low temperature without regard for orientation, and then re-embed the block in epoxy resin using heat polymerization. We have been doing this for years with Lowicryl HM20 polymerized at -50°C, re-embeding in epoxy resin at +60°C.

The sections have been probed with a wide variety of antibodies, which have all shown no detectable reduction in antigenicity (as compared with non-heat treated Lowicryl blocks).

Re-embedding in epoxy resin offers an excellent way of determining orientation at room temp. To keep the orientation during polymerization of epoxy resin, the Lowicryl blocks are first "glued" in place using only a small amount of resin in the flat embedding molds. Once the polymerization has been started by heating in the oven, fresh resin is added to the mold to make the correct block shape.

Another trick when working with Lowicryl and difficult-to-see embedded specimens is to "mark" the specimen location in the polymerized block (and also the perimeter of the resin block) with a small drop of nail varnish (polish?). The small drop of color can be used to locate the specimen after re-embedding in epoxy resin.

Sincerely,

Paul Webster, Ph.D.
House Research Institute
2100 W. 3rd St
Los Angeles
CA 90057, USA




-----Original Message-----
X-from: ehaller-at-health.usf.edu [mailto:ehaller-at-health.usf.edu]
Sent: Mon 1/14/2013 5:29 AM
To: Webster, Paul

Hi, Hong,

Is it possible to do a partial orientation on the samples, that is, to orient them in one direction, and polymerize them in the Lowicryl at the usual temperature, then cut the polymerized blocks with a saw, and Super-Glue the cut blocks onto other blocks to obtain your final orientation for sectioning?

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: hyi-at-emory.edu [hyi-at-emory.edu]
Sent: Sunday, January 13, 2013 12:52 PM
To: Haller, Edward

Dear Researchers:

I am doing a cryosubstitution run. The samples being processed need to
be positioned in a certain way for Lowicryl embedding. The process of
positioning samples takes time and needs to be done under a dissecting
scope, therefore it would be easier if it is done when the samples are at
a higher temperature (near zero), and subsequently embed/UV polymerize in
Lowicryl at the same temperature. Has anyone tried this? Is there
anything I need to be aware when UV polymerize Lowicryl at this
temperature? Thanks in advance.

Hong



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From: PhillipsT-at-missouri.edu
Date: Mon, 14 Jan 2013 10:01:51 -0600
Subject: [Microscopy] Lowicryl polymerization

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I agree with the sentiments of Paul and Debbie. I frequently embed my tissues in Lowicryl, LR Gold, and BMMA at 4 C and have excellent polymerization. I generally start with a flat-bottom BEEM capsule with a cap on it. I make no special effort to orient the tissue at the initial stage and Murphy's Law ensures that I get the wrong orientation even less often than chance might predict. I cut the blocks out of the BEEM capsule with a razor blade and inspect the surface with tissue in it under a stereoscope. I then mark the surface with a Sharpie and use a Dremel to trim away the plastic I don't want (i.e., where the Sharpie ink is). This generally results in a somewhat rectangle block that could be sectioned in a pinch but I usually insert this chunk of plastic back in a fresh BEEM capsule - the size of the block makes it easy to orient like I want and it can't easily shift in the capsule. I add a fresh label and resin and repolymerize. It costs me an extra day but I end up with a well-oriented block that is fully complete for inserting into the microtome chuck.

I find 4 C is sufficient for most antigens but if the immunolabeling doesn't work, I consider using a lower temperature embedding procedure. I list below three papers that discuss the effect of temperature on Lowicryl polymerization and/or immunolabeling. I also include a reference to a simple device I devised which allows tissues samples to be processed at sub-zero temperatures during dehydration, infiltration and polymerization. I built the device for less than $500 and it uses a conventional siphon-type carbon dioxide gas cylinder to maintain an aluminum block at temperatures as low as -35şC for over 15 hours/cylinder. Good luck. Tom

Armbruster BL, Garavito RM, Kellenberger E. 1983 Dehydration and embedding temperatures affect the antigenic specificity of tubulin and immunolabeling by the protein A-colloidal gold technique. J Histochem Cytochem 31(12):1380-1384.

Glauert A, Young RD. 1988. The control of temperature during polymerization of Lowicryl K4M: there is a low-temperature embedding method. J Microscopy 154(2):101-113.

Weibull C. 1987. Temperature rise in Lowicryl resins during polymerization by ultraviolet light. J Ultrastruct Molec Res 97:207-209.

Shoemaker, W., C. Hayes, and T.E. Phillips. 2003. A simple, low-cost device for processing and embedding tissues at sub-zero temperatures. Microscopy Research and Technique 62:262-266.


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: PWebster-at-hei.org [mailto:PWebster-at-hei.org]
Sent: Monday, January 14, 2013 9:33 AM
To: Phillips, Thomas E.


Dear All,

With Lowicryl, it is also possible to embed using UV illumination at low temperature without regard for orientation, and then re-embed the block in epoxy resin using heat polymerization. We have been doing this for years with Lowicryl HM20 polymerized at -50°C, re-embeding in epoxy resin at +60°C.

The sections have been probed with a wide variety of antibodies, which have all shown no detectable reduction in antigenicity (as compared with non-heat treated Lowicryl blocks).

Re-embedding in epoxy resin offers an excellent way of determining orientation at room temp. To keep the orientation during polymerization of epoxy resin, the Lowicryl blocks are first "glued" in place using only a small amount of resin in the flat embedding molds. Once the polymerization has been started by heating in the oven, fresh resin is added to the mold to make the correct block shape.

Another trick when working with Lowicryl and difficult-to-see embedded specimens is to "mark" the specimen location in the polymerized block (and also the perimeter of the resin block) with a small drop of nail varnish (polish?). The small drop of color can be used to locate the specimen after re-embedding in epoxy resin.

Sincerely,

Paul Webster, Ph.D.
House Research Institute
2100 W. 3rd St
Los Angeles
CA 90057, USA




-----Original Message-----
X-from: ehaller-at-health.usf.edu [mailto:ehaller-at-health.usf.edu]
Sent: Mon 1/14/2013 5:29 AM
To: Webster, Paul

Hi, Hong,

Is it possible to do a partial orientation on the samples, that is, to orient them in one direction, and polymerize them in the Lowicryl at the usual temperature, then cut the polymerized blocks with a saw, and Super-Glue the cut blocks onto other blocks to obtain your final orientation for sectioning?

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: hyi-at-emory.edu [hyi-at-emory.edu]
Sent: Sunday, January 13, 2013 12:52 PM
To: Haller, Edward

Dear Researchers:

I am doing a cryosubstitution run. The samples being processed need to be positioned in a certain way for Lowicryl embedding. The process of positioning samples takes time and needs to be done under a dissecting scope, therefore it would be easier if it is done when the samples are at a higher temperature (near zero), and subsequently embed/UV polymerize in Lowicryl at the same temperature. Has anyone tried this? Is there anything I need to be aware when UV polymerize Lowicryl at this temperature? Thanks in advance.

Hong



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From: tindallr-at-missouri.edu
Date: Mon, 14 Jan 2013 13:13:43 -0600
Subject: [Microscopy] Cryo tubing?

Contents Retrieved from Microscopy Listserver Archives
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Dear Collective,

Does anyone know of a source of flexible tubing for use with liquid nitrogen, outside diameter 6mm and inside diameter 4mm? Our cryoultramicrotome setup has tubing labeled as Festo PP-4, which the manufacturer says is rated down to -30 C and it cracks and breaks easily at LN2 temps. This dewar-to-cutting chamber hose can be easily repaired with the right tubing, but I'm not having any luck finding anything suitable in the right size.

Not optimistic, but eternally hopeful!

Cheers and thanks,
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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From: W.Muss-at-salk.at
Date: Mon, 14 Jan 2013 13:43:19 -0600
Subject: [Microscopy] Re(short): Cryo tubing?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

to: tindallr-at-missouri.edu

Good afternoon,
Dear Randy,
since here it is time to go home (8.40 p.m.) and your request was received some minute before leaving the lab hopefully very briefly:
my first thought was and my suggestion is (not very {inexpensive} I guess but most probably getting rid of broken hoses.)
I remember "old days" in the late 1970ies when I had a similar problem with tubing on an old cryo-ultramicrome...(Reichert OMU3 or was it an old Ultracut? with "cryochamber")
try Google Search {TEFLON} or PTFE tubing / tubes e.g. (no interest, no affiliation, just one out of the results found):

http://www.zeusinc.com/extrusionservices/materials/ptfe.aspx

PTFE Background Information
As the world's largest manufacturer of PTFE Tubing, Zeus offers a wide range of standard size products and precision custom PTFE Tubing. PTFE Tubing has become the gold standard in industries requiring the ultimate in lubricity, high temperature use, chemical resistance, biocompatibility, and precision extruded tolerances.
Key Properties
Very Lubricious - Lowest coefficient of friction of any polymer
Working temperature range 500° F (260° C) to -454° F (-270° C)
Chemically Resistant (all common solvents, acids and bases)
Chemically Inert
Low extractable
Excellent Dielectric Insulation Properties
For specific properties, see our Summary of Properties ....

and: http://www.ptfemart.com/ptfe-tubing.htm
it might be you'll find an appropriate out-/Inside diameter, for better thermic "shielding"(also not to get burned accidentally) one could wrap around foam-material ("thermo"-sleeving http://www.ptfemart.com/ptfe-tubing.htm material).

Good luck and best regards,

Wolfgang

Wolfgang MUSS
SALZBURG-AUSTRIA

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


Von: tindallr-at-missouri.edu [mailto:tindallr-at-missouri.edu]
Gesendet: Montag, 14. Jänner 2013 20:18
An: Muß Wolfgang
Betreff: [Microscopy] Cryo tubing?

----------------------------------------------------------------------------
The Microscopy ListServer -- CoSponsor: The Microscopy Society of America

Dear Collective,

Does anyone know of a source of flexible tubing for use with liquid nitrogen, outside diameter 6mm and inside diameter 4mm?
Our cryoultramicrotome setup has tubing labeled as Festo PP-4, which the manufacturer says is rated down to -30 C and it cracks and breaks easily at LN2 temps.

This dewar-to-cutting chamber hose can be easily repaired with the right tubing, but I'm not having any luck finding anything suitable in the right size.

Not optimistic, but eternally hopeful!

Cheers and thanks,
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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From: eric-miller-at-northwestern.edu
Date: Mon, 14 Jan 2013 14:20:27 -0600
Subject: [Microscopy] Hitachi SEM Instructions

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hey all, we've produced a few instructional videos here that go over how to use the Hitachi S-4800 and the S-3400. We've had some pretty good feedback on them and thought we would share them here in case anyone was interested.

Full 3400 instructions
http://youtu.be/BOUFKItQtHc

Quickie 3400 instructions
http://youtu.be/uQ8J2E3VzGk

Full 4800 instructions
http://youtu.be/52UKCY9fUR0



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



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From: Rosemary.White-at-csiro.au
Date: Mon, 14 Jan 2013 15:40:36 -0600
Subject: [Microscopy] Lowicryl polymerization

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Rather than re-embedding, if tissue is not correctly oriented for sectioning after polymerisation, we usually cut off the tip of the block with the specimen in such a way that we can superglue it, with tissue in correct orientation, onto a piece of perspex rod that's the right size for the microtome chuck. We get our workshop to cut the rod into the correct lengths and polish the ends. We get them to make a couple of hundred each time, it doesn't take them long and there's usually an apprentice who gets the job...

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
________________________________________
X-from: PhillipsT-at-missouri.edu [PhillipsT-at-missouri.edu]
Sent: Tuesday, 15 January 2013 3:06 a.m.
To: White, Rosemary (PI, Black Mountain)

I agree with the sentiments of Paul and Debbie. I frequently embed my tissues in Lowicryl, LR Gold, and BMMA at 4 C and have excellent polymerization. I generally start with a flat-bottom BEEM capsule with a cap on it. I make no special effort to orient the tissue at the initial stage and Murphy's Law ensures that I get the wrong orientation even less often than chance might predict. I cut the blocks out of the BEEM capsule with a razor blade and inspect the surface with tissue in it under a stereoscope. I then mark the surface with a Sharpie and use a Dremel to trim away the plastic I don't want (i.e., where the Sharpie ink is). This generally results in a somewhat rectangle block that could be sectioned in a pinch but I usually insert this chunk of plastic back in a fresh BEEM capsule - the size of the block makes it easy to orient like I want and it can't easily shift in the capsule. I add a fresh label and resin and repolymerize. It costs me an extra day but I end up wi!
th a well-oriented block that is fully complete for inserting into the microtome chuck.

I find 4 C is sufficient for most antigens but if the immunolabeling doesn't work, I consider using a lower temperature embedding procedure. I list below three papers that discuss the effect of temperature on Lowicryl polymerization and/or immunolabeling. I also include a reference to a simple device I devised which allows tissues samples to be processed at sub-zero temperatures during dehydration, infiltration and polymerization. I built the device for less than $500 and it uses a conventional siphon-type carbon dioxide gas cylinder to maintain an aluminum block at temperatures as low as -35şC for over 15 hours/cylinder. Good luck. Tom

Armbruster BL, Garavito RM, Kellenberger E. 1983 Dehydration and embedding temperatures affect the antigenic specificity of tubulin and immunolabeling by the protein A-colloidal gold technique. J Histochem Cytochem 31(12):1380-1384.

Glauert A, Young RD. 1988. The control of temperature during polymerization of Lowicryl K4M: there is a low-temperature embedding method. J Microscopy 154(2):101-113.

Weibull C. 1987. Temperature rise in Lowicryl resins during polymerization by ultraviolet light. J Ultrastruct Molec Res 97:207-209.

Shoemaker, W., C. Hayes, and T.E. Phillips. 2003. A simple, low-cost device for processing and embedding tissues at sub-zero temperatures. Microscopy Research and Technique 62:262-266.


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: PWebster-at-hei.org [mailto:PWebster-at-hei.org]
Sent: Monday, January 14, 2013 9:33 AM
To: Phillips, Thomas E.


Dear All,

With Lowicryl, it is also possible to embed using UV illumination at low temperature without regard for orientation, and then re-embed the block in epoxy resin using heat polymerization. We have been doing this for years with Lowicryl HM20 polymerized at -50°C, re-embeding in epoxy resin at +60°C.

The sections have been probed with a wide variety of antibodies, which have all shown no detectable reduction in antigenicity (as compared with non-heat treated Lowicryl blocks).

Re-embedding in epoxy resin offers an excellent way of determining orientation at room temp. To keep the orientation during polymerization of epoxy resin, the Lowicryl blocks are first "glued" in place using only a small amount of resin in the flat embedding molds. Once the polymerization has been started by heating in the oven, fresh resin is added to the mold to make the correct block shape.

Another trick when working with Lowicryl and difficult-to-see embedded specimens is to "mark" the specimen location in the polymerized block (and also the perimeter of the resin block) with a small drop of nail varnish (polish?). The small drop of color can be used to locate the specimen after re-embedding in epoxy resin.

Sincerely,

Paul Webster, Ph.D.
House Research Institute
2100 W. 3rd St
Los Angeles
CA 90057, USA




-----Original Message-----
X-from: ehaller-at-health.usf.edu [mailto:ehaller-at-health.usf.edu]
Sent: Mon 1/14/2013 5:29 AM
To: Webster, Paul

Hi, Hong,

Is it possible to do a partial orientation on the samples, that is, to orient them in one direction, and polymerize them in the Lowicryl at the usual temperature, then cut the polymerized blocks with a saw, and Super-Glue the cut blocks onto other blocks to obtain your final orientation for sectioning?

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: hyi-at-emory.edu [hyi-at-emory.edu]
Sent: Sunday, January 13, 2013 12:52 PM
To: Haller, Edward

Dear Researchers:

I am doing a cryosubstitution run. The samples being processed need to be positioned in a certain way for Lowicryl embedding. The process of positioning samples takes time and needs to be done under a dissecting scope, therefore it would be easier if it is done when the samples are at a higher temperature (near zero), and subsequently embed/UV polymerize in Lowicryl at the same temperature. Has anyone tried this? Is there anything I need to be aware when UV polymerize Lowicryl at this temperature? Thanks in advance.

Hong



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From: microscopylistserver-noreply-at-microscopy.com
Date: Mon, 14 Jan 2013 18:36:03 -0600
Subject: [Microscopy] viaWWW:RE:water-solvent swap

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Email: rleepenn-at-hotmail.com
Name: Lee Penn

Title-Subject: [Filtered] water-solvent swap

Message: In response to: Message: I have nanoparticles in what I suspect to be dirty water and
would like to transfer them into a clean solvent for drop-casting. Is it possible to evaporate the
water and then add the appropriate solvent or am I completely oblivious to some well known
nanoparticle in solution rule?
Thanks!

No - drying could change your particles irreversibly and cause substantial aggregation.

I recommend using dialysis against purified water.
If the things you want to remove from the suspension are dissolved, then the dialysis will result in
the diffusion of the dissolved species from the suspension to the surrounding purified water.

Check out the experimental section of this paper, and you will find details about how to perform
dialysis against purified water in order to remove dissolved species from a suspension of
nanoparticles and water.
Effect of Ionic Strength on the Kinetics of Crystal Growth by Oriented Aggregation, Nathan D.
Burrows, Christopher R. H. Hale, and R. Lee Penn (2012) Crystal Growth and Design, DOI:
10.1021/cg3004849 Article ASAP.

If you have a few more details about your nanoparticles, I'd be happy to chat about sample prep
options. We have worked with a range of nanoparticles in aqueous systems, ranging from very "dirty"
suspensions of natural materials in natural waters to nanoparticles harvested from aqueous reactors
to very clean systems that have been dialyzed against purified water.

Good luck and happy new year!



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From: microscopylistserver-noreply-at-microscopy.com
Date: Mon, 14 Jan 2013 18:37:09 -0600
Subject: [Microscopy] viaWWW:Reading beam current on ESEM Philips FEI XL30

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Email: gangqiu0-at-gmail.com
Name: Gqiu

Organization: Drexel University

Title-Subject: [Filtered] Reading beam current on ESEM Philips FEI XL30

Message: Hello all,

I am using a Philips FEI XL30 ESEM machine. I was wondering if it was possible to...
(1) measure the current that is striking the SAMPLE (not the filament).
(2) control the intensity of the beam current aside from changing the spot size (eg, specify a beam
with current of 1nA instead of spotsize 1)

My ultimate goal is to attain a beam with a current on the order of nano amps.

Any input will be appreciated,

Thank you
Gqiu

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From: forzaabbott-at-gmail.com
Date: Mon, 14 Jan 2013 20:21:58 -0600
Subject: [Microscopy] Re: viaWWW:Reading beam current on ESEM Philips FEI XL30

Contents Retrieved from Microscopy Listserver Archives
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Remembering back to when we did e-beam lithography on that microscope, there was a BNC connector on the door. We would plug our ammeter into this connector, and it seems that we had to disconnect a wire that grounded the stage inside the chamber. Disconnecting the wire would disable the touch sensor, so we had to be extra careful about not crashing the pole piece.

Jonathan Abbott

Sent from my mobile

On Jan 14, 2013, at 17:46, microscopylistserver-noreply-at-microscopy.com wrote:

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} Title-Subject: [Filtered] Reading beam current on ESEM Philips FEI XL30
}
} Message: Hello all,
}
} I am using a Philips FEI XL30 ESEM machine. I was wondering if it was possible to...
} (1) measure the current that is striking the SAMPLE (not the filament).
} (2) control the intensity of the beam current aside from changing the spot size (eg, specify a beam
} with current of 1nA instead of spotsize 1)
}
} My ultimate goal is to attain a beam with a current on the order of nano amps.
}
} Any input will be appreciated,
}
} Thank you
} Gqiu
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From: wesaia-at-iastate.edu
Date: Mon, 14 Jan 2013 20:32:07 -0600
Subject: [Microscopy] RE: viaWWW:Reading beam current on ESEM Philips FEI

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{Note that this is a reply to a web question. I copied them as well as the list server.}

Your request is quite normal, except for the part about doing it without changing the "spot size".

You will probably need a Faraday cup and a nano-ammeter to measure the current reaching your sample. I would like to think you could find such a meter without much trouble at Drexel. I don't know where you plug that into an XL30. I suppose it is on the front door, but others could say for sure. A Faraday cup may be as simple as small, deep hole drilled into a piece of carbon. If you want to get fancy, you can glue a used aperture over the top so that the electrons that enter the hole stay in the hole and get absorbed. In that case, sample current = beam current.

Now about "spot size" - I learned on SEMs that didn't have a "spot size" setting. They had condenser lens controls without any particular numbering. You adjusted the lens one way to choke the beam down for more resolution and the other way for more beam current. Of course, one of the side effects was that the size of the beam spot on the sample changed a little. I usually didn't worry about how much since I was working at lower magnifications (by today's standards). I choked it down for high resolution microscopy while maintaining a decent signal-to-noise ratio in my iamge. I increased it as necessary to get enough current for a decent count rate for x-ray analysis. The setting varied some depending on my needs and how much time I had. Of course, you can also change the objective aperture to minimize spot size for a given beam current. You'll have to experiment a little to see what gives you the sharpest image at 1 nA.

So again, what is your concern about the spot size? The three SEMs I work with now have spot sizes that range up to 7, 60, or 255. They are quite arbitrary and non-linear in their numbering. You should probably go through an exercise to see what your numbers correlate to in terms of incident beam size. I think your beam diameter will probably be no more than a few nm even at hefty currents. Hopefully you can focus better than many of our users so that you really do attain the optimum effect of that spot size.

My work tends to concentrate on microanalysis and the beam diameter is much smaller than the interaction volume diameter. I can safely ignore the beam diameter and just call it zero. Interaction volume is the main determinant of x-ray resolution. So now I spend a fair amount of time determining the interaction volume as a function of voltage in an effort to keep my interaction within the phase of interest while still exciting lines that I can resolve in the EDS spectra.

Of course, you may have a different issue at hand, maybe e-beam lithography, but I think you will still find that the interaction volume is more the determining factor. Set the condenser lens (spot size) where you need to at the voltage you need to.

Warren Straszheim
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Email: gangqiu0-at-gmail.com
Name: Gqiu

Organization: Drexel University

Title-Subject: [Filtered] Reading beam current on ESEM Philips FEI XL30

Message: Hello all,

I am using a Philips FEI XL30 ESEM machine. I was wondering if it was possible to...
(1) measure the current that is striking the SAMPLE (not the filament).
(2) control the intensity of the beam current aside from changing the spot size (eg, specify a beam
with current of 1nA instead of spotsize 1)

My ultimate goal is to attain a beam with a current on the order of nano amps.

Any input will be appreciated,

Thank you
Gqiu

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18, 37 -- Subject: RE: [Microscopy] viaWWW:Reading beam current on ESEM Philips FEI
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From: protrain-at-emcourses.com
Date: Tue, 15 Jan 2013 05:14:28 -0600
Subject: [Microscopy] RE: viaWWW:Reading beam current on ESEM Philips FEI XL30

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Hi

You have had a good deal of advice on this problem but may I address the
control of the beam current? There are three variables

1) Emission current from the gun - bias setting or filament position -
move the filament toward the cathode aperture for a higher current or
increase the emission current control.

2) The setting of the first condenser lens - stronger lens for a lower
current.

3) The size of the beam defining aperture - larger aperture for more
current.

X-from my knowledge there are hardly any, if any instruments that do directly
link the beam current with control of the condenser system. Whilst the
manufacturers will often guess the current they will be placing on the
specimen (their readout figure) this guess is only true for their idea of
where the above three variables are set; their guess is very much an
approximation! The only true indication of beam current is the Faraday Cup!

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

REPLY TO

Email: gangqiu0-at-gmail.com
Name: Gqiu

Organization: Drexel University

Title-Subject: [Filtered] Reading beam current on ESEM Philips FEI XL30

Message: Hello all,

I am using a Philips FEI XL30 ESEM machine. I was wondering if it was
possible to...
(1) measure the current that is striking the SAMPLE (not the filament).
(2) control the intensity of the beam current aside from changing the spot
size (eg, specify a beam with current of 1nA instead of spotsize 1)

My ultimate goal is to attain a beam with a current on the order of nano
amps.

Any input will be appreciated,

Thank you
Gqiu







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From: microscopylistserver-noreply-at-microscopy.com
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Subject: [Microscopy] viaWWW:Lehigh Microscopy School

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Message: There is still time to register for the 2013 Lehigh Microscopy School which will be held
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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 15 Jan 2013 19:48:10 -0600
Subject: [Microscopy] viaWWW:Confocal Workshop

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Title-Subject: [Filtered] Confocal Workshop

Message: The South Carolina EPSCoR/IDeA Program and the USC School of Medicine Instrumentation
Resource Facility are pleased to announce the 9th annual workshop on Basic Confocal Microscopy. The
Workshop will be held June 10-14 at the USC School of Medicine in Columbia, SC.

Workshop material is directed towards beginning and intermediate users of confocal microscopes and
involves a series of lectures (specimen preparation, labeling strategies, proper set-up of
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scanning and spinning disk confocal microscopes.

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Participants will have ample time for hands on use of the instruments during the workshop

Faculty will include Dr. Jay Jerome of Vanderbilt University, Dr. Ralph Albrecht of the University
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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 15 Jan 2013 19:49:04 -0600
Subject: [Microscopy] viaWWW:Post-doc available in electron microscopy

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Message: Quantum Non-Demolition Electron Microscope (QEM) Post-doctoral candidates are sought with
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will work with a small international team of researchers at MIT (K. Berggren and M. F. Yanik) and
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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 16 Jan 2013 08:11:01 -0600
Subject: [Microscopy] viaWWW:SEM chamber contamination

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Organization: نٍ شيَا خُوِ
ذُوْ ذِ جِ ثُوْ
宁夏大学

Title-Subject: [Filtered] SEM chamber contamination

Message: Dear List members,

Our laboratory recently purchased a “specially” commissioned SEM. The idea was that instrument was
as oil free as possible, so it was equipped with a TMP and a scroll pump. To our dismay, there is a
substantial contamination: the clearly visible infamous “dark squares”. They are currently
negotiating with the manufacturer but in the mean time we need to assess its quantity and type.
We are thinking of preparing some polished sample and collect a set of images at different
integration times and make a plot of darkness vs. time. The question is however, what would be an
acceptable “darkening” rate?

Does anyone know if there exists some sort of standardized protocol?

Can anyone recommend a procedure?

Some recommendations in dealing with the manufacturer are also welcome. Their first reaction was –
It is from your samples!!!. Fortunately, it was very easy to show this was not the case. Now they
are improvising all sorts of “cleanings”.

Thanks a lot
Mba


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From: les-at-zsgenetics.com
Date: Wed, 16 Jan 2013 08:56:26 -0600
Subject: [Microscopy] FW: viaWWW:SEM chamber contamination

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi Mba,
You should be able to scan an area for several minutes, without seeing
build-up. See the NIST paper (Evactron.com).

Short of a UHV solution, it is not surprising to have contamination in a SEM
or FIB. There are so many HC's diffusing out of crevices and off surfaces
from machined parts and plastics in the instrument, especially when it is
newer. If you have budget, an in-situ cleaning device like the GV-10x or
Evactron is extremely helpful.


Regards,
Larry Scipioni
ZS Genetics





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Organization: نٍ شيَا
خُوِ ذُوْ ذِ
جِ ثُوْ 宁夏大学

Title-Subject: [Filtered] SEM chamber contamination

Message: Dear List members,

Our laboratory recently purchased a “specially” commissioned SEM. The idea
was that instrument was as oil free as possible, so it was equipped with a
TMP and a scroll pump. To our dismay, there is a substantial contamination:
the clearly visible infamous “dark squares”. They are currently
negotiating with the manufacturer but in the mean time we need to assess its
quantity and type.
We are thinking of preparing some polished sample and collect a set of
images at different integration times and make a plot of darkness vs. time.
The question is however, what would be an acceptable “darkening” rate?

Does anyone know if there exists some sort of standardized protocol?

Can anyone recommend a procedure?

Some recommendations in dealing with the manufacturer are also welcome.
Their first reaction was – It is from your samples!!!. Fortunately, it was
very easy to show this was not the case. Now they are improvising all sorts
of “cleanings”.

Thanks a lot
Mba


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From: ian-at-acutance.co.uk
Date: Wed, 16 Jan 2013 10:25:01 -0600
Subject: [Microscopy] viaWWW:SEM chamber contamination

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

For a protocol you might look at:

Conru and LaBerge, Oil Contamination with SEM Operated in Spot Scan Mode. J
Phys E Sci Instr 8(2): 136-138, 1975.



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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Organization: نٍ شيَا
خُوِ
ذُوْ ذِ جِ
ثُوْ
宁夏大学

Title-Subject: [Filtered] SEM chamber contamination

Message: Dear List members,

Our laboratory recently purchased a “specially” commissioned SEM. The idea
was that instrument was
as oil free as possible, so it was equipped with a TMP and a scroll pump. To
our dismay, there is a
substantial contamination: the clearly visible infamous “dark squares”.
They are currently
negotiating with the manufacturer but in the mean time we need to assess its
quantity and type.
We are thinking of preparing some polished sample and collect a set of
images at different
integration times and make a plot of darkness vs. time. The question is
however, what would be an
acceptable “darkening” rate?

Does anyone know if there exists some sort of standardized protocol?

Can anyone recommend a procedure?

Some recommendations in dealing with the manufacturer are also welcome.
Their first reaction was –
It is from your samples!!!. Fortunately, it was very easy to show this was
not the case. Now they
are improvising all sorts of “cleanings”.

Thanks a lot
Mba


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Listserver Email Form V - 20120416
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From noreply-at-onlinebrusselsguide.com Wed Jan 16 10:17:51 2013
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Mba

Do contact XEI Scientific Inc., in Redwood, Ca. They sell devices which plug
into SEMs to decontaminate them of hydrocarbons. You can find them at
http://www.evactron.com/

Best Regards

Ian

Ian Holton
Acutance Scientific


-----Original Message-----
X-from: ph2-at-sprynet.com [mailto:ph2-at-sprynet.com]
Sent: 16 January 2013 16:14
To: ian-at-acutance.co.uk

For a protocol you might look at:

Conru and LaBerge, Oil Contamination with SEM Operated in Spot Scan Mode. J
Phys E Sci Instr 8(2): 136-138, 1975.



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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Organization: نٍ شيَا
خُوِ
ذُوْ ذِ جِ
ثُوْ
宁夏大学

Title-Subject: [Filtered] SEM chamber contamination

Message: Dear List members,

Our laboratory recently purchased a “specially” commissioned SEM. The idea
was that instrument was
as oil free as possible, so it was equipped with a TMP and a scroll pump. To
our dismay, there is a
substantial contamination: the clearly visible infamous “dark squares”.
They are currently
negotiating with the manufacturer but in the mean time we need to assess its
quantity and type.
We are thinking of preparing some polished sample and collect a set of
images at different
integration times and make a plot of darkness vs. time. The question is
however, what would be an
acceptable “darkening” rate?

Does anyone know if there exists some sort of standardized protocol?

Can anyone recommend a procedure?

Some recommendations in dealing with the manufacturer are also welcome.
Their first reaction was –
It is from your samples!!!. Fortunately, it was very easy to show this was
not the case. Now they
are improvising all sorts of “cleanings”.

Thanks a lot
Mba


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From: FMonson-at-wcupa.edu
Date: Wed, 16 Jan 2013 11:39:51 -0600
Subject: [Microscopy] viaWWW:water-solvent swap

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

What is the 'dirt'?
If it sediments then decant the less dirty supernate.
If what's decanted has dirt bigger than the nano-s, then filter - with forethought.
If the filtrate has dissolved material then use dialysis that won't pass the nano-s to reduce the volume wholistically and simultaneously concentrating the nano-s.
You might also use differential centrifugation.

All of the above depends on the size of the nano-s, and how they behave, AND whether you actually know they are present.

Hope this helps,

Fred Monson

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
West Chester University of Pennsylvania
Schmucker Science South - Room SSS-024
MailDrop: Geology-Astronomy
750 South Church Street
West Chester, PA, 19383
610-738-0437
fmonson-at-wcupa.edu

HomePage: http://cmirt.wcupa.edu (with link to instrument Scheduler)



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Name: Marissa

Organization: LBNL

Title-Subject: [Filtered] water-solvent swap

Message: I have nanoparticles in what I suspect to be dirty water and would like to transfer them into a clean solvent for drop-casting. Is it possible to evaporate the water and then add the appropriate solvent or am I completely oblivious to some well known nanoparticle in solution rule?

Thanks!

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From: kenconverse-at-qualityimages.biz
Date: Wed, 16 Jan 2013 12:44:02 -0600
Subject: [Microscopy] viaWWW:SEM chamber contamination

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Mba,
I remember reading something in an old Kurt Lesker catalogue stating that if
you use a Turbo pump like a diffusion pump, with isolation valving so that
the pump can run continuously, it will be very clean. On the other hand, if
you run it the way many do, stopping it and bringing it to atmosphere every
time you vent the chamber, you will get oils creeping up through the turbo
pump and contaminating the chamber. This explained to me why SEMs that I
worked on where the DP had been replaced with a turbo were very clean and
Amrays (in particular) that had turbos and a single vent valve were so
terribly dirty.

Is your turbo in a system with roughing, backing and high vacuum isolation
valves, or just a single vent valve? If your turbo is mag-lev, then it is
most likely isolated and runs all the time due to the limited life of the
crash bearings. Otherwise, you might have the "simple" system, and it will
most likely be dirty.

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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Organization: نٍ شيَا
خُوِ
ذُوْ ذِ جِ
ثُوْ
宁夏大学

Title-Subject: [Filtered] SEM chamber contamination

Message: Dear List members,

Our laboratory recently purchased a “specially” commissioned SEM. The idea
was that instrument was
as oil free as possible, so it was equipped with a TMP and a scroll pump. To
our dismay, there is a
substantial contamination: the clearly visible infamous “dark squares”.
They are currently
negotiating with the manufacturer but in the mean time we need to assess its
quantity and type.
We are thinking of preparing some polished sample and collect a set of
images at different
integration times and make a plot of darkness vs. time. The question is
however, what would be an
acceptable “darkening” rate?

Does anyone know if there exists some sort of standardized protocol?

Can anyone recommend a procedure?

Some recommendations in dealing with the manufacturer are also welcome.
Their first reaction was –
It is from your samples!!!. Fortunately, it was very easy to show this was
not the case. Now they
are improvising all sorts of “cleanings”.

Thanks a lot
Mba


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From: kenconverse-at-qualityimages.biz
Date: Wed, 16 Jan 2013 13:11:07 -0600
Subject: [Microscopy] RE: viaWWW:Reading beam current on ESEM Philips FEI

Contents Retrieved from Microscopy Listserver Archives
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I believe that there is a basic limiting factor and that is electron current
density. This is pretty much determined by the source, and with thermionic
sources, when you increase the emission current you increase the area from
which electrons are emitted, all else being equal, you have a larger spot.

When you increase your condenser lens current, you get a smaller spot, but
with many fewer electrons. The same is true when you change final aperture
size.

In the end, there will be essentially a fixed spot size for a given beam
current when everything is focused. If the spot size you want is larger
than the nominal spot size for the current you need, you can just defocus
the beam. If you need a smaller spot for a given current, you need a
different electron source that will yield a higher current density.

Given all the variables in setting up a beam (kV, emission current,
condenser current, "objective" current, working distance, aperture sizes),
the only way to have a known current is to measure it using a Faraday cup
and pico-ammeter, as explained in other posts. You may be able to establish
some "standard settings" that will readily put you in the ball park, but if
the current is critical, it must be measured. Anyone doing quantitative
x-ray analysis or small scale ebeam lithography will confirm this.

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: Tuesday, January 15, 2013 6:17 AM
To: kenconverse-at-qualityimages.biz

Hi

You have had a good deal of advice on this problem but may I address the
control of the beam current? There are three variables

1) Emission current from the gun - bias setting or filament position -
move the filament toward the cathode aperture for a higher current or
increase the emission current control.

2) The setting of the first condenser lens - stronger lens for a lower
current.

3) The size of the beam defining aperture - larger aperture for more
current.

X-from my knowledge there are hardly any, if any instruments that do
directly
link the beam current with control of the condenser system. Whilst the
manufacturers will often guess the current they will be placing on the
specimen (their readout figure) this guess is only true for their idea of
where the above three variables are set; their guess is very much an
approximation! The only true indication of beam current is the Faraday Cup!

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

REPLY TO

Email: gangqiu0-at-gmail.com
Name: Gqiu

Organization: Drexel University

Title-Subject: [Filtered] Reading beam current on ESEM Philips FEI XL30

Message: Hello all,

I am using a Philips FEI XL30 ESEM machine. I was wondering if it was
possible to...
(1) measure the current that is striking the SAMPLE (not the filament).
(2) control the intensity of the beam current aside from changing the spot
size (eg, specify a beam with current of 1nA instead of spotsize 1)

My ultimate goal is to attain a beam with a current on the order of nano
amps.

Any input will be appreciated,

Thank you
Gqiu







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From: stefan.diller-at-t-online.de
Date: Wed, 16 Jan 2013 13:32:42 -0600
Subject: [Microscopy] LaB6 cathode - double contours in SEM image

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear All,
today I mounted a new Denka LaB6 cathode first time and after firing it up got double contours on the image details.
Can anybody give me a hint if there is some basic problem with the geometry in the Wehnelt, e.g. cathode too far away or too near?
I mounted it as I normally mount Kimball LaB6 cathodes, centered very carefully, with a distance of the tip of ca. 0.15mm from the
UPPER (facing the Anode) Wehnelt plane.
It seems like there are two sources of electrons going out of the crystall. It might also come from the more "brick-like" shape of
the LaB6 cristal, in comparison with the round shape of a Kimball tip... When I heat the cathode up some notches more (I am now
nearly at the end of the scale), it`s still the same. I can post a link with images tomorrow, but today I missed taking some...
I put the same cathode in my Philips TEM EM420 and the image at 80 KV is perfect, also the cathode image during heating-up is as
it should be (in the TEM...).
Shall I first try and go nearer with the Wehnelt? Current at 30KV and medium Bias setting is now ca. 40 uA.

Any ideas?

Thanks,
Stefan


--


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Websites:
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Anfahrt: http://Mail.map24.com/Stefan.Diller
-----------------------------------------------------

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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 16 Jan 2013 19:59:27 -0600
Subject: [Microscopy] viaWWW:ISI-SS40 documentation

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Email: gregc-at-physics.ucsb.edu
Name: Greg Courville

Organization: University of California, Santa Barbara

Title-Subject: [Filtered] ISI-SS40 documentation?

Message: Hello all,

We've just acquired an ISI-SS40 scanning electron microscope, with no documentation other than some
photocopied instructions and a manual for a different model (SX40). We contacted the service company
whose sticker we found on the unit, but they don't even have the docs for this model around any
more. If anyone has any documentation on this unit (in particular, schematics and/or a service
manual) that they wouldn't mind parting with or scanning, we'd love to hear from you and would be
willing to reimburse you as appropriate. Any other information on this series (we understand the
SS-60 and SS-160 are substantially similar instruments) would be greatly appreciated as well.

Best regards
--
Greg Courville

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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 16 Jan 2013 20:00:07 -0600
Subject: [Microscopy] Fwd: [Filtered] MicroscopyListserverviaWWW:

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Title-Subject: [Filtered] Graphene/scotch tape preparation

Message: Hello again. Is someone willing to share or recommend literature on a method for making a
graphene TEM sample using the scotch tape trick? Does this film need washing (HCl sol?) or plasma
cleaning to guarantee the adhesive is fully removed?

Thank you!

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From: philip.oshel-at-cmich.edu
Date: Fri, 18 Jan 2013 12:22:08 -0600
Subject: [Microscopy] Ask-A-Microscopist HR SEM, kV, and surface topography

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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 - Manuela Anstoetz
Email - manuela.anstoetz-at-scu.edu.au
ORGANIZATION - Southern Cross University
EDUCATION - Graduate College
LOCATION - Lismore, NSW, Australia
SUBJECT_OF_QUESTION - HRSEM and Ultra-HRSEM
QUESTION - Hello,
I would like to know if for HRSEM and Ultra-HRSEM Microscopy the premise
of low accelerating voltage gives more surface information and high
accelerating voltage delivers more sub-surface information is cancelled?
I have seen 30 kV being used for surface topology of a metallic whisker
growing from a thin film and feel confused about that. I looked up the
Microscope used (Hitachi SU5500) and discovered that it has a range for
Vacc from 0.5 keV to 30keV. I would have used 5keV or so to determine
surface morphology. Can you help clarify, please?
Thank you kindly,
Manuela


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From: protrain-at-emcourses.com
Date: Sun, 20 Jan 2013 04:43:34 -0600
Subject: [Microscopy] RE: Ask-A-Microscopist HR SEM, kV, and surface topography

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Hi Manuela

First let me say that unfortunately not all the SEM results that you see in
publications are indicative of good microscopy! So much of what I see
suggests to me that the editors accept publications from those who know very
little about SEM! Now let's get down to your question.

No matter which SEM you use, no matter how clever its imaging system, the
basic physics of signal generation apply.

a) Secondary electrons are liberated from near the surface of the
material no matter what accelerating voltage is used.
b) Backscattered electrons are liberated from a volume below the
surface, the higher the accelerating voltage the deeper the depth from which
they may be generated.
c) Backscattered electrons striking parts of the chamber also produce
these signals. The BSE being converted into multiple secondary electrons
that may then contribute to the image. These converted BSE may be very
useful, are less effected by charge, therefore they often make a difficult
specimen usable. The problem is the converted BSE still carry the BSE data,
so the SE collected actually depict the sub surface information.

So to answer your question the lower the accelerating voltage the better if
you wish to see the true surface of a specimen. Lowering the accelerating
voltage reduces the depth of penetration allowing the SE signal to dominate.
Raising the accelerating voltage creates a larger volume from which BSE may
be produced. In this case the BSE themselves, or the converted BSE, may
dominate the information subduing the surface information.

You correctly note that the Hitachi have provided you with a very wide range
of accelerating voltages indicating that there is a very difficult and
critical decision to be made when you select your operating voltage.
Unfortunately 85% of operators around the world use the same accelerating
voltage no matter what type of specimen they are viewing. As a guide if you
wish to see the true surface of a specimen, you are correct, you should
operate at less than 5kV. However in a number of applications we may use
the collection of signals from different depths by changing the accelerating
voltage, "sectioning by kV", this technique often provides a mass of useful
information.

TTL(through the lens) detection is a very efficient method for collecting
SE, but if too many of the converted BSE are excluded charge may become a
major problem. Thus most knowledgeable operators will vary the WD with a
TTL system to vary the amount of converted BSE used to subdue charge. The
alternative is to work with a TTL system at very low accelerating voltages,
certainly less than 2kV, and to rely upon the superior collection efficiency
of the system to allow small spot sizes to be used, to minimise the probe
current thus minimise the charge. I am often working with clients at 1 to
1.5kV up to 150,000X in this type of system. We do not need to coat if we
carefully balance accelerating voltage with WD to make the impossible
possible.

Remember, there will be a particular accelerating voltage that will provide
more information than any other. How do we find this voltage? Well
microscopists are scientists and a scientist are meant to experiment!

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

****************************************************************************
************
realname - Manuela Anstoetz
Email - manuela.anstoetz-at-scu.edu.au
ORGANIZATION - Southern Cross University EDUCATION - Graduate College
LOCATION - Lismore, NSW, Australia SUBJECT_OF_QUESTION - HRSEM and
Ultra-HRSEM QUESTION - Hello, I would like to know if for HRSEM and
Ultra-HRSEM Microscopy the premise of low accelerating voltage gives more
surface information and high accelerating voltage delivers more sub-surface
information is cancelled?
I have seen 30 kV being used for surface topology of a metallic whisker
growing from a thin film and feel confused about that. I looked up the
Microscope used (Hitachi SU5500) and discovered that it has a range for Vacc
from 0.5 keV to 30keV. I would have used 5keV or so to determine surface
morphology. Can you help clarify, please?
Thank you kindly,
Manuela





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From: dsherman-at-purdue.edu
Date: Sun, 20 Jan 2013 12:33:30 -0600
Subject: [Microscopy] Ask-A-Microscopist HR SEM, kV, and surface

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Manuela,

I agree entirely with Steve's comments below, but want to extend the
conversation a bit in a slightly different direction. You are the perfect
example of the importance of not just teaching but "education" to make a
good microscopist.

The easy way out is to sit down at a microscope and have someone show you
what knobs to turn or buttons to push or screen functions to activate to
get an image. It is also the fastest way to get users for your
instruments. Some of these users will question and gradually educate
themselves, but the majority will not. However, in my opinion, this is a
real disservice to not only to you, the user, but to others that use the
instrumentation for two reasons:

1) As Steve so ably explained below, you need to understand enough about
the principles behind SEM (and other forms of microscopy) so as to use the
systems to your advantage "in an educated way" to get the most out of the
instrument and your sample. Understanding the hows and whys of basic
operation will make you able to adapt not only to different samples and
the information needed from that sample, but also to different microscopes
as you advance your career. A good solid course in microscopy principles
is not a waste of time. It actually will speed up good data accumulation
down the line with more in-depth understanding of your samples in the
process.

2) Understanding how an instrument works and recognizing when a system is
not working to its full potential will help not only you but all the other
users of the instrument. You will often be able to recognize potential
problems, alert the appropriate people, and thus get the problems resolved
before they become serious. Also, responsible use of major equipment goes
hand-in-hand with understanding the system and also your limitations in
dealing with that system, so that you do not unknowingly or
unintentionally compromise the instrument.

It comes down to EDUCATION not just training if we want to develop, or be,
more than just instrument technicians, but truly competent microscopists.
Unfortunately microscopy courses are to often eliminated or diluted down
due to pressures to produce NOW not after completing a course as well as
less emphasis on education by faculty/supervisors/facility directors &
managers as they are confronted with these pressures.

Debby


Debra Sherman, Founder and CSO
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





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From: protrain-at-emcourses.com
Date: Sun, 20 Jan 2013 13:23:31 -0600
Subject: Re: [Microscopy] RE: Ask-A-Microscopist HR SEM, kV, and surface

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi

Back again to add to the comments Debby made.

A good friend of mine from Australia has this headline in his laboratory "Do
YOU do science or do YOU do tradition." Think about it?

Steve

-----Original Message-----
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Sent: 20 January 2013 18:33
To: protrain-at-emcourses.com; message to: MSA list;
manuela.anstoetz-at-scu.edu.au




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23, 24 -- Subject: RE: [Microscopy] RE: Ask-A-Microscopist HR SEM, kV, and surface topography-education!
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From: kenconverse-at-qualityimages.biz
Date: Sun, 20 Jan 2013 15:31:25 -0600
Subject: Re: [Microscopy] RE: Ask-A-Microscopist HR SEM, kV, and surface

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

To amplify what Debbie and Steve are saying:

I have always loved SEMs because they are a signal generator and a signal
processor, not a microscope. If you tell me what you want to see, I can
probably show it to you, then we need to have a long talk about what is
real. (What, you found argon in another sample?).

The world NEEDS microscopists! Anyone can take a pretty picture with the
new instruments, but are you sure of what the image is showing 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: protrain-at-emcourses.com [mailto:protrain-at-emcourses.com]
Sent: Sunday, January 20, 2013 2:26 PM
To: kenconverse-at-qualityimages.biz

Hi

Back again to add to the comments Debby made.

A good friend of mine from Australia has this headline in his laboratory "Do
YOU do science or do YOU do tradition." Think about it?

Steve

-----Original Message-----
X-from: Sherman, Debra [mailto:dsherman-at-purdue.edu]
Sent: 20 January 2013 18:33
To: protrain-at-emcourses.com; message to: MSA list;
manuela.anstoetz-at-scu.edu.au




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surface topography-education!
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From: microscopylistserver-noreply-at-microscopy.com
Date: Mon, 21 Jan 2013 08:51:53 -0600
Subject: [Microscopy] viaWWW:NESM February, 28th Meeting @ Boston University

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Marissa Libbee asked about "nanoparticles in [suspected] dirty water". I
don't have an answer to her question but want to share my own findings and
ask others for comments.

I'm studying nanoparticles in aqueous solution. The particles are in the
range 1-5 nm diameter. My task is to image them by AFM and to measure the
height of many individual particles. The sample preparation procedure is
basically to apply a droplet of the suspension to freshly cleaved mica,
rinse and dry. The AFM images are made in air at room temperature using
TappingMode (intermittent contact) and obtain nice images. However, blank
samples prepared from the ultrapure water used as diluent and as rinse water
also show some particles.
I have tried more than 3 different samples of ultrapure water produced using
Milli-Q water purification systems at 3 different labs and gotten similar
results.
I wonder whether anyone else has had similar experiences working with water
and mica.

regards,
Don
=============================================
Don Chernoff, Ph.D., President
Advanced Surface Microscopy, Inc.; 3250 N. Post Rd., Suite 120;
Indianapolis IN 46226 USA
E-Mail: donc-at-asmicro.com www.asmicro.com
Voice: 317-895-5630 Toll free: 800-374-8557 (in USA & Canada)
Fax: 317-895-5652
[business activities since 1990: analytical services in AFM, AFM probes,
consulting, training, calibration and test specimens,
calibration and measurement software, used NanoScope equipment.]
=============================================
----- Original Message -----
From: FMonson-at-wcupa.edu
To: donc-at-asmicro.com
Sent: Wednesday, January 16, 2013 12:42 PM
Subject: [a] [Microscopy] RE: viaWWW:water-solvent swap





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What is the 'dirt'?
If it sediments then decant the less dirty supernate.
If what's decanted has dirt bigger than the nano-s, then filter - with
forethought.
If the filtrate has dissolved material then use dialysis that won't pass
the nano-s to reduce the volume wholistically and simultaneously
concentrating the nano-s.
You might also use differential centrifugation.

All of the above depends on the size of the nano-s, and how they behave,
AND whether you actually know they are present.

Hope this helps,

Fred Monson

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
West Chester University of Pennsylvania
Schmucker Science South - Room SSS-024
MailDrop: Geology-Astronomy
750 South Church Street
West Chester, PA, 19383
610-738-0437
fmonson-at-wcupa.edu

HomePage: http://cmirt.wcupa.edu (with link to instrument Scheduler)



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Subject: [Microscopy] viaWWW:water-solvent swap




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Email: mlibbee-at-gmail.com
Name: Marissa

Organization: LBNL

Title-Subject: [Filtered] water-solvent swap

Message: I have nanoparticles in what I suspect to be dirty water and
would like to transfer them into a clean solvent for drop-casting. Is it
possible to evaporate the water and then add the appropriate solvent or am I
completely oblivious to some well known nanoparticle in solution rule?

Thanks!

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Email: mzugravu-at-nesmicroscopy.org
Name: Monica Viorica Zugravu

Organization: Harvard University, Center for Nanoscale Systems

Title-Subject: [Filtered] NESM February, 28th Meeting -at- Boston University

Message: Greetings Fellow Microscopists,

This is your friendly Save the Date reminder for the NESM February Meeting hosted by Boston
University on February 28. The meeting will consist of a BU Photonics Center tour, a buffet dinner,
and two technical talks. Stay tuned for detailed information about the February Meeting in the
coming weeks. You can also keep up with NESM activities by checking us out on the web at
nesmicroscopy.org and on Facebook (https://www.facebook.com/NESMicroscopy) and Twitter
(https://twitter.com/#!/NESMicroscopy).

Cheers,
NESM Board


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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 22 Jan 2013 07:15:44 -0600
Subject: [Microscopy] viaWWW:Open Position: Research Assistant Electron Microscopy

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Email: poper-at-nbacc.net
Name: Robert Pope

Organization: BNBI/NBACC

Title-Subject: [Filtered] Open Position: Research Assistant Electron Microscopy

Message: JOB TITLE: Research Assistant, Electron Microscopy
Company:Battelle National Biodefense Institute www.bnbi.org

PRIMARY FUNCTION

Provides biological sample preparation support for Scanning Electron Microscopy (SEM) and
Transmission Electron Microscopy (TEM) and performs Scanning and Transmission Microscopy of samples
to support the identification and characterization of biological agents and animal tissues.

MAJOR RESPONSIBILITIES:

Prepares samples containing biological agents for SEM and TEM analysis.
Operates and maintains NBACC Scanning Electron Microscope.
Operates and maintains NBACC Transmission Electron Microscope.
Operates and maintains NBACC Electron Microscopy ancillary equipment.
Supports the operation of the NBACC Core Electron Microscopy capability.
Works with a team of scientists collaborating to develop, manage, and analyze research and to
conduct bioforensic analyses critical to the mission of the NBACC.
Maintains records, analyzes technical and programmatic data and prepares analytical reports.
Presents results in written and oral formats.
Attends specialized training and reviews safety manuals and SOPS in order to maintain a safe
workplace environment.
Performs work with select agents in BSL2, BSL3 and BSL4 laboratories with appropriate training.
Identifies departures from the Quality Management System (QMS) and initiates actions to investigate
and prevent such occurrences.
Performs other assigned duties.

MINIMUM REQUIRED QUALIFICATIONS:
Bachelors degree (or equivalent) preferably in a biological science field. Experience in general
laboratory, biological, and electron microscopy techniques preferred.
Experience in specific techniques including embedding, fixing, sectioning, and staining of
biological samples for scanning and transmission electron microscopy desired.
Experience in the interpretation of SEM and TEM images for a variety of biological samples desirable.
General skills in light microscopy, phase contrast microscopy and fluorescent microscopy desirable.
Familiarity with imaging of BSL-2/3/4 viruses and bacteria, and familiarity with the relevant
safety, biosurety, decontamination, and quality assurance guidelines desirable.
Proficiency of oral and written communications essential.
Exemplary organizational skills with a proven track record of working effectively both independently
and as a team player.
Knowledge or experience with biocontainment facilities and procedures, laboratory safety, biosurety,
and decontamination desirable.
Knowledge or experience with Quality Assurance (QA) programs such as ISO 9001 or ISO 17025 and/or
GMP/GLP work environments desirable.
Must be a citizen of the United States.
Must be able to obtain an Interim secret clearance, leading to an eventual top secret clearance,
Suitability for Department of Homeland Security (DHS), a favorable adjudication of the Department of
Justice (DoJ) for select agents access, and enrollment in the Biological and Personnel Reliability
Program.
Current clearance(s) desirable.
Required to enroll in the Special Immunizations Program (SIP).

Apply at http://www.bnbi.org/careers.html

https://home.eease.com/recruit2/?id=3620041&t=1


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From: tina-at-pbrc.hawaii.edu
Date: Tue, 22 Jan 2013 18:23:08 -0600
Subject: [Microscopy] Surface replica of wet sample for SEM

Contents Retrieved from Microscopy Listserver Archives
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I have a client who needs to make a replica of a wet surface
(gelatin-like) for our field emission SEM. I've seen a reference to
polyvinylsiloxane or Xantropen Blue. Before I run around trying to locate
this stuff, does anyone have a recommendation for any material, plus a
source? Hints and tips?

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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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 23 Jan 2013 07:38:08 -0600
Subject: [Microscopy] viaWWW:Want CCD camera for Gatan Image Filter

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Hi

Two (euro) cents more to Steve, Debby and Ken very good general comments :

As far I know, the Hitachi SU5500 is a HR SEM working in a TEM like
configuration, with the sample (cut to a smale size) in the OL, between
the upper and the lower pole piece, at WD = 0.
If the sample is a thin sample, and if the SEM is fited with a
transmited electrons detector (note the two "if"), it works in a way
similar to a STEM, and in that case the best resolution conditions will
be at high beam energy. Note that I say "resolution" conditions, not
"surface informations" conditions.
Look carefully at the indications on the picture or what is said in the
paper : which detector was used, SE (upper) or STEM ? What about the
sample preparation ? Is the picture a side or cross section like view,
or a top view ? Maybe the explanation of the choice of 30 keV is there.
Maybe too, that the conditions were choosen for STEM mode, but a picture
in SE-TTL was made, and gave THE information needed to answer THE question.
I'll hope that someone using a SU5500 know the way to use a SEM, but...

Hope it helps

--

J. Faerber
IPCMS-DSI
Institut de Physique et Chimie des Matériaux de Strasbourg
Département 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: crozier-at-asu.edu
Name: Peter Crozier

Organization: Arizona State University

Title-Subject: [Filtered] Want CCD camera for Gatan Image Filter

Message: Dear Colleagues,
The CCD camera on the back of our Gatan Imaging Filter (GIF) has failed and is apparently too old to
be repaired according to Gatan. The system is installed on an FEI Tecnai F20 TEM and it is the main
detector for our EELS capability. The electron optics for the GIF is fine it is just the detector
that has failed. We are interested in hearing from anyone that may have a suggestion on how to
repair the system. Ideally we would like to acquire a used CCD camera that is compatible with our GIF.
The model number of our current camera is 794GIF.3K3BP.33. It is a “SI033 Site CCD”. Any 794GIF in
the serial# range of 01102000 up to 04020200 will have this CCD.

We would also consider a complete replacement of the GIF with a used GIF or used PEELS system (we
donÂ’t perform image filtering on our Tecnai).

If any of you are able to help us secure this repair please contact me. We appreciate any
assistance and suggestions you may have.

Thanks

Peter Crozier
Associate Professor
Arizona State University


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From: philip.oshel-at-cmich.edu
Date: Wed, 23 Jan 2013 11:02:10 -0600
Subject: [Microscopy] Ask-A-Microscopist metal coating depth into pores

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***************************************************************************************
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realname - Soya
Email - sgamsey-at-hotmail.com
EDUCATION - Graduate College
QUESTION - Is it possible to coat a porous sample (~80um pore size) in a
way that allows deposition of the coating material (e.g. Pt) into the pores?

--


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From: protrain-at-emcourses.com
Date: Wed, 23 Jan 2013 11:14:56 -0600
Subject: [Microscopy] Ask-A-Microscopist metal coating depth into pores

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi

If you are considering sputter coating there is a method I have used but not
for pores as small as you have.

Place the sample in the sputter coater and pump it to attain the best
possible vacuum. Do not bleed gas into the system but try to force it to
coat; automated systems may not let you do this old systems will. The
theory is that with very little gas you have more straight line deposition,
the only chance of putting the coat inside the pores.

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: philip.oshel-at-cmich.edu [mailto:philip.oshel-at-cmich.edu]
Sent: 23 January 2013 17:03
To: protrain-at-emcourses.com

--





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From: jkabel-at-mail.ubc.ca
Date: Wed, 23 Jan 2013 13:36:11 -0600
Subject: [Microscopy] Re: Ask-A-Microscopist metal coating depth into pores

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi Soya,

Sputter and evaporative coating methods are line of sight to the
source. This means if you want to coat into a porous surface, you have
to play games with the geometry. The most common way would be with a
sample spinner, wherein the sample is doubly rotated on an eccentric
path, usually while tilted. This will only allow you to coat those
pores with line of sight to the source from any possible orientation.

-Jacob

On 13-01-23 09:12 AM, philip.oshel-at-cmich.edu wrote:
}
}
} ----------------------------------------------------------------------------
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} ----------------------------------------------------------------------------
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} 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 - Soya
} Email - sgamsey-at-hotmail.com
} EDUCATION - Graduate College
} QUESTION - Is it possible to coat a porous sample (~80um pore size) in a
} way that allows deposition of the coating material (e.g. Pt) into the pores?
}

--
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
http://www.mtrl.ubc.ca/research/eml/


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From: stefan.diller-at-t-online.de
Date: Wed, 23 Jan 2013 14:27:35 -0600
Subject: [Microscopy] LKB Histo Knifemaker 2078 manual needed

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear All,
anybody out there who can send me the manual of the LKB Histo Knifemaker 2078 in PDF?

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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From: kraftpiano-at-gmail.com
Date: Wed, 23 Jan 2013 15:02:31 -0600
Subject: [Microscopy] Ask-A-Microscopist metal coating depth into pores

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I'm going to defer to others for the specifics on this, but wouldn'
tan Osmium tetra oxide treatment work for getting into the porous
surface cavities? I know it's not exactly the most environmentally
friendly solution, but would that work better than the sputter coating
methods?

--Justin A. Kraft

On Wed, Jan 23, 2013 at 2:41 PM, {jkabel-at-mail.ubc.ca} 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 Soya,
}
} Sputter and evaporative coating methods are line of sight to the
} source. This means if you want to coat into a porous surface, you have
} to play games with the geometry. The most common way would be with a
} sample spinner, wherein the sample is doubly rotated on an eccentric
} path, usually while tilted. This will only allow you to coat those
} pores with line of sight to the source from any possible orientation.
}
} -Jacob
}
} On 13-01-23 09:12 AM, philip.oshel-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
} } ----------------------------------------------------------------------------
} }
} } ***************************************************************************************
} } 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 - Soya
} } Email - sgamsey-at-hotmail.com
} } EDUCATION - Graduate College
} } QUESTION - Is it possible to coat a porous sample (~80um pore size) in a
} } way that allows deposition of the coating material (e.g. Pt) into the pores?
} }
}
} --
} 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
} http://www.mtrl.ubc.ca/research/eml/
}
}
} ==============================Original Headers==============================
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--
"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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From: protrain-at-emcourses.com
Date: Wed, 23 Jan 2013 15:57:09 -0600
Subject: [Microscopy] Ask-A-Microscopist metal coating depth into pores

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi Soya,

Time of coat depends upon your coater, perhaps try 1 minute coats. Tilt 45
degs in one direction for the first coat, then 45 degs in the opposite
direction for the second. For the final coat move to the best vacuum as
described.

In my experience the higher vacuum coat is important because the object of
sputter coating is to cause the metal to be deviated away from line of site
by the gas. The better vacuum increases the mean free path of the metal and
that is the only way to penetrate holes.

An interesting experiment for the doubters is to take a nut (as in nuts and
bolts) about 1.16inch hole diameter and see which technique puts metal
through the hole.

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: Soya Gamsey [mailto:sgamsey-at-hotmail.com]
Sent: 23 January 2013 18:39
To: protrain-at-emcourses.com

--








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28, 23 -- References: {201301231702.r0NH2q4D000747-at-ns.microscopy.com} {006701cdf98d$2b4d3ec0$81e7bc40$-at-com} {BLU0-SMTP310F1F58E254F98C07787CCB8150-at-phx.gbl}
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From: rice-at-nysbc.org
Date: Wed, 23 Jan 2013 17:01:38 -0600
Subject: [Microscopy] Staff position in the EM Facility at New York Structural Biology

Contents Retrieved from Microscopy Listserver Archives
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Staff position in the EM Facility at New York Structural Biology Center


The New York Structural Biology Center (NYSBC) seeks an experienced
electron microscopist to join the staff of its cryo-EM facility. The
NYSBC is a shared center that supports state-of-the-art research in
cryo-EM, NMR, and X-ray crystallography. Cryo-EM facilities includes
four transmission electron microscopes, one dual-beam FIB/SEM, and
related support equipment for grid preparation, high pressure freezing,
and sectioning. In general, projects focus on 3D reconstruction of
biological assemblies, such as the atomic structure of membrane
proteins, subunit organization of macromolecular complexes, and cellular
anatomy of various tissues. The individual will carry out experiments in
support of collaborative projects with affiliated investigators,
including EM sample preparation, data collection, and image processing.
Day to day duties include setup and maintenance of electron microscopes,
training and assisting users from NYSBC's nine Member Institutions, and
record keeping. The individual should be capable of multitasking and
enjoy working with other people. Good communication skills are
essential. Qualified applicants should send a curriculum vitae and names
of three references to David Stokes (_stokes-at-nysbc.org
{mailto:stokes-at-nysbc.org} _). Salary and job title will be commensurate
with experience. A Ph. D. is desirable but not essential for this
position. The position is currently open and applications will be
reviewed continuously until the position is filled.

--
William J. Rice, Ph.D.
Senior Scientist, EM Facility Manager
New York Structural Biology Center
89 Convent Avenue, NY, NY 10027



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From: dsherman-at-purdue.edu
Date: Wed, 23 Jan 2013 19:40:31 -0600
Subject: [Microscopy] Ask-A-Microscopist metal coating depth into

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Soya,

This is a perfect sample for low vacuum or low voltage imaging if you have
those capabilities.

Also, you can try coating with carbon using a vacuum evaporator. In this
case the carbon is projected down on the sample with minimal direction
change. Thus chances are better to get some down into the pores. However,
do keep in mind that you need to evaporate in a good vacuum to get the
finest coating. This along with metal shadowing to more effectively cover
the surface of your sample may be adequate to minimize charge accumulation.

Debby


Debra Sherman, Founder & CSO
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 1/23/13 4:58 PM, "protrain-at-emcourses.com" {protrain-at-emcourses.com}
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From: nizets2-at-yahoo.com
Date: Thu, 24 Jan 2013 01:07:49 -0600
Subject: [Microscopy] dehydration for histopathology in mouse

Contents Retrieved from Microscopy Listserver Archives
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Dear colleagues,

I am planning a mouse study which has to be performed by a private company. In their protocol to prepare the mouse intestine for histopathological investigation, they describe their protocol like that:

"In brief, colons (one part of the longitudinally divided colons) are fixed as “Swiss-rolls” overnight in 10% neutral buffered formalin. Then colons are transferred directly into 70% ethanol, rolled, processed(Technicon Autotechnicon Mono Tissue Processor.This automated tissue processor sequentially submerses tissue specimens into solutions of fixative, dehydrant, clearant and paraffin wax at timed intervals)and embedded into paraffin blocks. Sections of the size of 5 µm will be cut for haematoxylin andeosin staining. "
 
As a cell biology this dehydration looks really minimalistic. Personally I would add more step but I suppose that the protocols used by pathologists for routine analysis have been adapted for higher throughput.
The purpose is to investigate modifications of the histology of the tissue (rough changes), no special staining here.
 
I would like to have the feedback of pathologists to know if this is usual to reduce the dehydration to one step of ethanol.
Many thanks in advance.
 
Stephane


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From: W.Muss-at-salk.at
Date: Thu, 24 Jan 2013 03:29:46 -0600
Subject: [Microscopy] Re: dehydration for histopathology in mouse

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Good morning,
dear Stephane,

the description of process by the company’s information says:

“in brief“... and further “…sequentially submerses tissue specimens into solutions of fixative, dehydrant, clearant and paraffin wax at timed intervals)”.
Therefore this does not necessarily mean that there is only “one step of dehydrating agent = ethanol”!

So you and we do not know which protocol the private company for the automated tissue processor exactly will follow.
You will have to ask for the detailed, complete processing manual including all steps until paraffin blocking (see below) , but I am sure, you would be able also (if you are paying client) to “order” a processing schedule YOU would like to be implemented for “your” special tissue.

In routine pathological tissue processing – as I understand and as I know from our Histology Lab - a routine schedule (times depending on the material to be processed, here you will get the times for our so called “fat containing tissue”-processing schedule - ) will be
(further evaluation of resulting paraffin blocks –as you know- by means of EM/TEM usually not recommended but can be done by “re-processing” and “re-embedding” into resin):

- Fixation in 10% neutral buffered formalin = 4% buffered Formaldehyde solution (commercially available, usually PO4-buffered):
since most of tissues sent to our Histo-Pathology Lab have been immersed in fixative already for transport, after macroscopy/grossing the small tissue objects are put into the plastic specimen cassettes for subsequent processing into paraffin. Usually the thickness of a sample at least in one dimension does not exceed 5 mm, but most of such sliced specimens are in the range of 2 -4 mm thickness (occasionally for larger biopsies applying also pressure or vacuum using special “fixator” devices) up to overnight (5-10 hrs) with or without increased temperature up to 35°/40°C for special tissues application)

after grossing: storage of the tissue containing cassettes in fixative usually -at- RT until automated processing:

Depending on the processor type (modern ones are sucking solutions from a concentrated stock solution bottle and dilute to the concentrations you would like / want to use automatically) you have to program an appropriate processing schedule: e.g. duration of each step / iteration / temperature and concentrations.

So in our Histo-Lab there is the following schedule (for “fat-containing” tissues) with regard to the automated processor (a Sakira Tissue-Tek) used:

- at least 1/2 hr -at- RT or elevated T 4% buffered formaldehyde solution
(- a washing step in buffer would be desirable but usually is not done to save TAT)
- Transfer from fixative directly into (70% ) 76% Ethanol (or Isopropanol) (1-2x, each at least 1hr) 1 hr
- Ethanol 80% : 1 hr
- Ethanol 96%: 3 x 1 hr
- Ethanol 100%: 2 x 1 hr
- clearant(s): varying from lab to lab: in our Histolab specimens are transferred within the tissue processor from the 3rd Ethanol 100%-bath directly into xylene (3 x 1 hr each). NB: other Labs use Xylene substituents (like Histoclear, Limonene, or similar).
Paraffin wax: Paraffin (melting point 65°C)4 x 45 mins each.

After Paraffin-impregnation, tissues are blocked on the plastic cassette (“Ausgiessen”, cf. “modular embedding centers” consisting of paraffin dispensing unit, warming trays for cassettes and cooling plate), cooled and then are ready for sectioning.

Disclaimer: mentioning of names of companies or chemicals only for information. No financial interest, no affiliation.

Hope this is helpful for you,

Wolfgang


Wolfgang MUSS
EM-Lab
Pathology
SALK-LKH (Gen. Hospital)
SALZBURG, AUSTRIA



} -----UrsprĂĽngliche Nachricht-----
} Von: nizets2-at-yahoo.com [mailto:nizets2-at-yahoo.com]
} Gesendet: Donnerstag, 24. Jänner 2013 08:11
} An: MuĂź Wolfgang
} Betreff: [Microscopy] dehydration for histopathology in mouse
}
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}
} Dear colleagues,
}
} I am planning a mouse study which has to be performed by a private
} company. In their protocol to prepare the mouse intestine for
} histopathological investigation, they describe their protocol like
} that:
}
} "In brief, colons (one part of the longitudinally divided colons) are
} fixed as “Swiss-rolls” overnight in 10% neutral buffered formalin. Then
} colons are transferred directly into 70% ethanol, rolled, processed
} (Technicon Autotechnicon Mono Tissue Processor. This automated tissue
} processor sequentially submerses tissue specimens into solutions of
} fixative, dehydrant, clearant and paraffin wax at timed intervals) and
} embedded into paraffin blocks. Sections of the size of 5 µm will be cut
} for haematoxylin andeosin staining. "
}
} As a cell biology this dehydration looks really minimalistic.
} Personally I would add more step but I suppose that the protocols used
} by pathologists for routine analysis have been adapted for higher
} throughput.
} The purpose is to investigate modifications of the histology of the
} tissue (rough changes), no special staining here.
}
} I would like to have the feedback of pathologists to know if this is
} usual to reduce the dehydration to one step of ethanol.
} Many thanks in advance.
}
} Stephane
}
}
} ==============================Original
} Headers==============================
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From: ben.micklem-at-pharm.ox.ac.uk
Date: Thu, 24 Jan 2013 03:34:34 -0600
Subject: [Microscopy] Re: dehydration for histopathology in mouse

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Dear Stephane,

I'm not familiar with the Autotechnicon Mono, but from Googling, it seems
that the tissue processor does multiple stages of dehydration.

I'm not sure why there is the preceding step of directly placing into 70%
ethanol­ just guessing, but maybe if this is left out the rolls of tissue
are not tightly/compactly enough rolled after the automated dehydration?

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 24/01/2013 07:14, "nizets2-at-yahoo.com" {nizets2-at-yahoo.com} wrote:

}
} Dear colleagues,
}
} I am planning a mouse study which has to be performed by a private
} company. In their protocol to prepare the mouse intestine for
} histopathological investigation, they describe their protocol like that:
}
} "In brief, colons (one part of the longitudinally divided colons) are
} fixed as łSwiss-rolls˛ overnight in 10% neutral buffered formalin. Then
} colons are transferred directly into 70% ethanol, rolled,
} processed(Technicon Autotechnicon Mono Tissue Processor.This automated
} tissue processor sequentially submerses tissue specimens into solutions
} of fixative, dehydrant, clearant and paraffin wax at timed intervals)and
} embedded into paraffin blocks. Sections of the size of 5 µm will be cut
} for haematoxylin andeosin staining. "
}
} As a cell biology this dehydration looks really minimalistic. Personally
} I would add more step but I suppose that the protocols used by
} pathologists for routine analysis have been adapted for higher throughput.
} The purpose is to investigate modifications of the histology of the
} tissue (rough changes), no special staining here.
}
} I would like to have the feedback of pathologists to know if this is
} usual to reduce the dehydration to one step of ethanol.
} Many thanks in advance.
}
} Stephane



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From: microscopylistserver-noreply-at-microscopy.com
Date: Thu, 24 Jan 2013 08:27:48 -0600
Subject: [Microscopy] viaWWW:Digital camera for JEOL CX100

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Email: yamawaki-at-stanford.edu
Name: Ruth Yamawaki

Organization: Stanford University

Title-Subject: [Filtered] Digital camera for JEOL CX100

Message: We are finally entering the digital age and are looking for a digital camera for our JEOL
CX100.

Please send me any tips/comments/service on your digital camera that can help us make an informed
decision.

Thank you,
Ruth Yamawaki
Stanford University
Department of Comparative Medicine
Stanford CA 94305
650 723-3457

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From: microscopylistserver-noreply-at-microscopy.com
Date: Thu, 24 Jan 2013 08:29:02 -0600
Subject: [Microscopy] viaWWW:Keep FEGTEM column clean

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Email: z.zhou-at-lboro.ac.uk
Name: Zhaoxia Zhou

Organization: Loughborough University

Title-Subject: [Filtered] Keep FEGTEM column clean

Message: We have a FEI Tecnai F20 TEM. Just installed, column nice and clean. STEM/HAADF mode hours
not seen the scan box/lines at all. Minimal contamination.

However, IÂ’ve got requests to look at some sol-gel prepared nanoparticles. Would heating the samples
(nanoparticles on holey carbon film) in an oven ~150°C for an hour be an effective way to prevent
possible contamination to the TEM? Are there other methods colleagues would like to recommend? What
are the usual procedures to keep an FEGTEM column clean with a wide range of types of samples?

Kind regards,
Zhaoxia Zhou

Dr Z Zhou
Experimental Officer
Loughborough Materials Characterisation Centre (LMCC)
Department of Materials
Loughborough University
Leicestershire
LE11 3TU
UK


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From: jinguo.wang-at-utdallas.edu
Date: Thu, 24 Jan 2013 09:45:09 -0600
Subject: [Microscopy] TEM/STEM postdoc position opening

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The University of Texas at Dallas has an opening for a postdoc position on the TEM/STEM study on nanostructured materials.

A Ph. D. in Materials Science or related physical science with experience on Transmission Electron Microscopy; knowledge of nanomaterials materials (especially, graphene, BN, MoS2, multilayer GaN/InN, etc.) and their structure-property relationships. Extensive experience with advanced electron microscopy documented by high-quality publications; hands-on experience on HRTEM and STEM with analytical techniques (EELS and EDS).

Interested candidates should provide a cover letter, CV, and a list of references to jinguo.wang-at-utdallas.edu. Applications will be evaluated as received.

Thanks

Jinguo Wang, Ph.D
Mail Station RL10
Department of Materials Science
University of Texas at Dallas
800 W. Campbell Road
Richardson, TX 75080-3021

jinguo.wang-at-utdallas.edu
972-883-6928




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From: ian-at-acutance.co.uk
Date: Thu, 24 Jan 2013 10:02:24 -0600
Subject: [Microscopy] viaWWW:Keep FEGTEM column clean

Contents Retrieved from Microscopy Listserver Archives
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Zhaoxia,

XEI Scientific Inc have a "TEM Wand" which replaces the sample-probe and
cleans up the sample region in a TEM by injecting Oxygen radicals into the
pole-region space.

Best Regards

Ian

Ian Holton
Acutance Scientific


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Email: z.zhou-at-lboro.ac.uk
Name: Zhaoxia Zhou

Organization: Loughborough University

Title-Subject: [Filtered] Keep FEGTEM column clean

Message: We have a FEI Tecnai F20 TEM. Just installed, column nice and
clean. STEM/HAADF mode hours not seen the scan box/lines at all. Minimal
contamination.

However, IÂ’ve got requests to look at some sol-gel prepared nanoparticles.
Would heating the samples (nanoparticles on holey carbon film) in an oven
~150°C for an hour be an effective way to prevent possible contamination to
the TEM? Are there other methods colleagues would like to recommend? What
are the usual procedures to keep an FEGTEM column clean with a wide range of
types of samples?

Kind regards,
Zhaoxia Zhou

Dr Z Zhou
Experimental Officer
Loughborough Materials Characterisation Centre (LMCC) Department of
Materials Loughborough University Leicestershire
LE11 3TU
UK


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From: tindallr-at-missouri.edu
Date: Thu, 24 Jan 2013 10:59:48 -0600
Subject: [Microscopy] TEM Neg staining: bacteria grown on carbon films

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear Collective,

Somewhere in the dim mists of memory seems to lurk a notion that I once saw a protocol for growing bacteria directly on carbon-coated TEM grids. So far, I've been unable to locate any trace of it, so either I have lost it or I am losing it. Does anybody know of techniques for doing this?

Thanks for any help you can give me.

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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From: tindallr-at-missouri.edu
Date: Thu, 24 Jan 2013 11:08:21 -0600
Subject: [Microscopy] TEM neg stain: bacteria, part 2

Contents Retrieved from Microscopy Listserver Archives
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Me, again. I forgot the second part of my question about culturing bacteria---are there ways of removing excess extracellular polysaccharides from bacterial cultures in order to cut down on the background garbage in negative stain? Our client is going to try a "protocol gradient", if I understand him correctly (I please ignorance on this), but we're coming up short on other ideas.

Thanks again!

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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From: FMonson-at-wcupa.edu
Date: Thu, 24 Jan 2013 12:36:55 -0600
Subject: [Microscopy] TEM neg stain: bacteria, part 2

Contents Retrieved from Microscopy Listserver Archives
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Long, long ago, when using the Periodic Acid Schiff (PAS) Rx on 'fresh', paraffin-embedded (that means well cooked, dewaxed, dehydrated and fixed (factually denatured)) tissue sections (all of that done to something like liver so it would look like the picture Maximov drew with his colored inks and published in one of his many histology HARDBOUND books on histology*) those of us who required a 'control' section were EXPECTED to acquire it by processing a second section with freshly collected saliva either collected solely by the efforts of one or, of course, better statistically if acquired thru contributions of a large, representative group of saliva producers. The result was more magenta in the test and less in the control. Pure science!

Using that method, one NEVER considered a gradient of anything in a protocol. However, one could, even in that dark age, purchase a pure example of an appropriate enzyme from Worthington.

* Histology has almost disappeared as a subject that synthesizes, anatomy, physiology, development, and often some pathology. My guess is that it requires sufficient knowledge of anatomy, microscopy, and those other subjects to build a curriculum of obligatory study - something that does not ring the school bells these days. Except for one exception, written for pathologists, normal histology now comes with a soft cover just to prove that it is not going to last much longer.

Cheers,

Fred Monson

Frederick C. Monson, PhD
Technical Director
Center for Microanalysis and Imaging, Research and Training (CMIRT)
West Chester University of Pennsylvania
Schmucker Science South - Room SSS-024
MailDrop: Geology-Astronomy
750 South Church Street
West Chester, PA, 19383
610-738-0437
fmonson-at-wcupa.edu

HomePage: http://cmirt.wcupa.edu (with link to instrument Scheduler)

-----Original Message-----
X-from: tindallr-at-missouri.edu [mailto:tindallr-at-missouri.edu]
Sent: Thursday, January 24, 2013 12:14 PM
To: Monson, Frederick

Me, again. I forgot the second part of my question about culturing bacteria---are there ways of removing excess extracellular polysaccharides from bacterial cultures in order to cut down on the background garbage in negative stain? Our client is going to try a "protocol gradient", if I understand him correctly (I please ignorance on this), but we're coming up short on other ideas.

Thanks again!

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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From: microscopylistserver-noreply-at-microscopy.com
Date: Thu, 24 Jan 2013 18:40:25 -0600
Subject: [Microscopy] viaWWW:TEM/STEM postdoc/visiting scientist position opening

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Email: tprozoro-at-ameslab.gov
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Organization: Ames Laboratory

Title-Subject: [Filtered] TEM/STEM postdoc/visiting scientist position opening

Message: Emergent Atomic and Magnetic NanoStructures group at the Ames Laboratory is seeking a
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From: W.Muss-at-salk.at
Date: Mon, 28 Jan 2013 05:23:00 -0600
Subject: [Microscopy] Re: Prep for platelets for TEM

Contents Retrieved from Microscopy Listserver Archives
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Dear Randy,

since I have not seen any reaction or reply to your post (1) with
the exception of Fred Monson's post on TEM Neg staining(2)

I would like to point you cautiously to the following papers

---------------------------------------------------------------------
cf.
www.bss.phy.cam.ac.uk/~es10009/documents/thomson_scanning2011.pdf

= SCANNING VOL. 33, 59-68 (2011)
& Wiley Periodicals, Inc.
=====Imaging Internal Features of Whole, Unfixed Bacteria=======
NICHOLAS M. THOMSON1, KEVIN CHANNON1, NOOR AZLIN MOKHTAR2,
LECH STANIEWICZ1, RANJANA RAI3,IPSITA ROY3, SHUN SATO4,
TAKEHARU TSUGE4, ATHENE M. DONALD1, DAVID SUMMERS2, AND
EASAN SIVANIAH1
1Cavendish Laboratory, University of Cambridge, Cambridge, UK
2Department of Genetics, University of Cambridge, Cambridge, UK
3Department of Molecular and Applied Biology, University of Westminster,
London, UK
4Department of Innovative and Engineered Materials, Tokyo Institute of
Technology, Yokohama, Japan
DOI 10.1002/sca.20221 Published online 22 February 2011 in
Wiley Online Library (wileyonlinelibrary.com)
--------------------------------------------------------------------
Most important with regard to your initial quest:

==== In situ multiple sampling of attached bacteria for scanning
and transmission electron microscopy.=====
Bozzola JJ, Johnson MC, Shechmeister IL.
Stain Technol. 1973 Nov;48(6):317-25. No abstract available.
PMID: 4129036 [PubMed - indexed for MEDLINE]
(If you need a / this .pdf, please just request it with your reply)

--------------------------------------------------------------------
and cf. (free access) -at-
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC246670/pdf/jbacter00340-0326.pdf
J Bacteriol. 1974 April; 118(1): 304-311. PMCID: PMC246670
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC246670/
===== Morphological Study of Streptococcus mutans and
Two Extracellular Polysaccharide Mutants =========

by M. C. Johnson, J. J. Bozzola, and I. L. Shechmeister

ex methods (it is more/less the same method as used in the
Stain Technol. 1973 Nov;48(6):317-25-article):

Electron microscopy.
A technique was developed which allowed electron micrographs to be taken of bacteria as they appeared when grown on a hard surface in a liquid medium containing sucrose. In this procedure,
a sterilized microscope slide,
a cover slip, and

{ {a cover slip with Formvar-carbon-coated grids were placed in a sterile plastic petri dish} } .

Sucrose broth was added, inoculated with the desired organism, and incubated at 35 C for a predetermined time. The above components were then treated as follows. The slide and plain cover slip were removed and gently washed two to three times in double-distilled water; the slide was air-dried for three to four days, and the cover slip was fixed overight in 3% gluteraldehyde and buffered to pH 6.0 with Veronal acetate.

The dried slide was processed for replicas (17), and the

cover slip was rinsed thoroughly in double-distilled water, air- or freeze-dried, coated with 40 to 50 nm of palladium-gold (60 to 40 alloy), and examined in a Kent Cambridge Mark II Stereoscan operating at an accelerating voltage of 30 kV and at a 450 angle tilt.

{ {The cover slip with attached grids} }
was dipped three times in double-distilled water, and the grids were removed to a filter paper and stained with 2% phosphotungstic acid at pH 5.5.

The bacteria on the bottom of the petri dish were then embedded in situ as follows: after a gentle wash with Veronal acetate, the sample was fixed for 24 h in Veronal acetate-buffered 1% OSO4 (16), dehydrated, and gradually infiltrated with Epon at 0.5 h-intervals by three changes of 3:1, 1:1, 1:3 (vol/vol) mixtures of absolute ethanol-Epon. After three more changes of pure Epon, a thin layer of resin was poured onto the petri dish and polymerized at 60 C for 48 h. The hardened plastic was allowed to cool for 24 h, after which the Epon layer was stripped off, sectioned, and examined with a Hitachi HU-11A transmission electron microscope at an accelerating voltage of 50 kV.

---------------------------------------------------------------------------
and last but not least (old too, but perhaps also interesting, and
Reinhard Rachel, Regensburg,
who is also on the MSA-List can/will comment on that):

J Bacteriol. 2006 October; 188(19): 6915-6923.
doi: 10.1128/JB.00527-06
PMCID: PMC1595509
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC1595509/
Flagella of Pyrococcus furiosus: Multifunctional Organelles,
Made for Swimming, Adhesion to Various Surfaces, and Cell-Cell Contacts
Daniela J. Näther,1 Reinhard Rachel,1 Gerhard Wanner,2 and Reinhard Wirth1
Adherence studies: growth on carbon-coated gold grids for TEM.
Methods to study growth of microorganisms
{ { directly on carbon-coated gold grids used for TEM } } have been
developed in our labs ([39] notice: + Ref.: see below).
Gold grids were placed in small Teflon holders in serum bottles
containing anaerobic medium. For TEM cells were fixed with 2.5%
(final concentration) glutaraldehyde for 30 min at room temperature.
In the case of cell or flagellum suspensions, a drop was placed on
a carbon-coated 200-mesh copper grid (Plano, Wetzlar, Germany).
The samples were either unidirectionally shadowed with Pt/C at
15° (CFE 50; Cressington Ltd., Watford, United Kingdom) or
negatively stained for 1 min with 2% uranyl acetate.
All TEM micrographs were recorded using a slow-scan charge-coupled
device camera (TEM 1000; TVIPS-Tietz, Gauting, Germany) attached
to a CM 12 transmission electron microscope (FEI, Eindhoven,
The Netherlands) operated at 120 keV.

[39]. Rieger, G., R. Rachel, R. Herrmann, and K. O. Stetter. 1995.
Ultrastructure of the hyperthermophilic archaeon Pyrodictium abyssi.
J. Struct. Biol. 115:78-87.
unfortunately this article does not show up in PubMed and
did not result as a Google-search finding.
------------------------------------------------------------------------

There also one could consider to use "lacey Support-Film" Grids:

http://www.tedpella.com/supflm_html/suptfilm.htm
7. Lacey Support Films - NetMeshT Grids:

Lacey Support Film

A lacey network support film. The holes in the lacey support film
vary in size from less than a quarter micron to more than 10 microns
making them ideal for any type of specimen. Lacey support films are
extremely strong and withstand vigorous specimen preparation treatment.

The specimen material is supported by the film network but lies across
or protrudes into the holes of the mesh. This allows high definition
imaging without the effects of underlying support material. Lacey films
can be used for specimens ranging from large crystals and other
particulate material to virus particles. Smaller particles,
such as viruses or bacteria,
tend to adhere around the inner edges of the holes, an ideal situation
for high resolution microscopy. Lacey films are also ideal for selected
area electron diffraction imaging. We offer three types of lacey film:

http://www.tedpella.com/supflm_html/suptfilm.htm
Support Film Grids,

Substrate Application Guide
B= Best; G= Good Alternative; -= Not Suitable
------------------------------------------------------------------------x
Substrate Formvar Formvar Silicon Silicon Carbon Carbon
Application Only Stabilized Monoxide Monoxide Type-A Type-B
with Carbon on Formvar on Type-A

Bacterial -NO- G B B B B
Suspensions
------------------------------------------------------------------------x

x-----------------------------------------------------------------------z
Substrate Pure Is Lacey Film
Application Carbon suitable for
Film this application?
Bacterial
Suspensions B Yes
x-----------------------------------------------------------------------z


Hope this helps you at least a little bit to get started,
sorry again (apologizing) for the longness of my posting
best regards and
to all of you listers
a happy and joyful weekend,

Wolfgang MUSS PhD
Salzburg - Austria




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} An: Muß Wolfgang
} Betreff: [Microscopy] TEM Neg staining(1): bacteria grown on carbon
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} Dear Collective,
}
} Somewhere in the dim mists of memory seems to lurk a notion that I once
} saw a protocol for growing bacteria directly on carbon-coated TEM
} grids.
}
} So far, I've been unable to locate any trace of it, so either I have
} lost it or I am losing it.
} Does anybody know of techniques for doing this?
}
} Thanks for any help you can give me.
}
} Cheers,
} Randy
}
} Randy Tindall
} Senior Research Specialist
} Electron Microscopy Core Facility
} W125 Veterinary Medicine
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Dear Judy,
unfortunately I have not seen any response to your post via MSA-Listserver. You might have gotten some reply's on this, so apologize if I am bothering with my posting.
The procedure of collecting blood cells for TEM most frequently is the preparation of a so called "Buffy Coat" (in the literature or in www- recipes I have found several misspellings for this term).

So, just to get started (there may exist hundreds of other or more sophisticated protocols):

1) Usually one will collect platelets from peripheral blood (the more ml the better - knowing this to be an ethical question)
2) Use e. g. EDTA-peripheral blood (filled into 'Monovettes' or similar pre-treated tubes with EDTA for blood sampling). There are out also Heparin-layered tubes. Avoid/ DON'T use collecting systems containing small glass globules! You'll never get a {buffy coat} .
2) You'll need a preparatory centrifuge, for preparing a "buffy coat" a centrifuge with a swing-out rotor is suited best (there are some small centrifuges which have implemented not only fix angle rotor, but swing out rotor as well, if you own a (cooled) bench top centrifuge you for sure are on the right side)
NB: cf. e.g. http://www.hettichlab.com/appc/content_manager/page.php?ID=160391&dbc=fa06a43d4d0ce718387d80170d8decd7
See: e.g. Small centrifuges: EBA 270, EBA 21; or BenchTop centrifuges...+/- all types, spec. rotors are parts to order)
(Disclaimer: there are a lot of centrifuge manufacturers. This URL only for a rough information. No financial interest, no affiliation, just having been satisfied user of one of their products over many years)
3) Before centrifugation topple tubes carefully / with caution some times (3-4 times)
4) make sure you'll have at least 2 tubes with the same weight (total tube+blood)
5) centrifuge blood for 10 min -at- 1000 g (the g-force will depend on radius of the rotor as well as the velocity, so you'll have to check this with the centrifuge's nomogramm which normally is enclosed with the centrifuge manual)
6) don't use a "break-function" of the centrifuge, that means, after centrifugation let swing out the rotor without disturbing the slowing of rotation until the rotor stands still

7) Carefully/cautiously take out the vials - put them into a fitting (tube)rack.
8) You have prepared in advance your fixative for TEM (there are a lot of possibilities, as you know), use at RT
9) In the Monovettes/tubes you should have gotten a "Buffy coat", that means layers of different dense "fluids" (from above to bottom), containing:
- Plasma/Serum (somehow yellowish-brown, depending on lipids in serum
- a thin layer of somehow whitish substrate (= platelet fraction)
- a thin layer of red/reddish substrate ( on top of that layer you'll find lymphocytes, on ground of that layer you 'll have spinned down neutrophils etc.
- bottom (+/less thick): RBC-Red blood Cells.
CAVE: don't use rotation forces higher than maximally recommended for the plastic tubes! 1.000 g is enough!

Take care when handling this because it is all fluid and the layer formation can be disturbed by any breath of a 'kick'.

Fixation of "Buffy Coat":
10) Gently/carefully open the tubes, suck off Plasma by means of a long pipette tip (slowly and positioning the tip in the middle of the tube. Control removal of serum carefully. Leave only a small layer of serum over the first opaque looking layer (taking care not to suck off from that layer with the pipette).
11) Gently release your TEM- fixative solution ONLY DROP BY DROP from another pipette tip which you place closely to the inner wall of the tube. Observe initial mixing of plasma remnants with the fixative. NO WHIRLING should be produced with that initial step of fixation.
(Cave: the thicker the left plasma layer will be the less you'll face problems with whirling, on the other hand you'll need longer time to get initially fixed the cells below (i.e. platelets, lymphocytes, neutrophils, etc).
After some minute add more drops of fixative slowly (at the end, fill up the tube)
If you use initially a primary fixative (e. g. 0.5 % FA + 1.5% GA buffered n 0.1M PO4-(or sodium cacodylate) buffer, let stand at RT for at least 15 -30 min UNDISTURBED on its own. Afterwards you can suck off the primary fixative and fill in cautiously your main fixative (e.g. 3-4% GA buffered in 0.13M PO4 = isotonic for blood or 0.1M cacodylate buffer).
12: let stand for either at least 3 hrs at RT or try out also "mild" MW-fixation in the tubes.

Collection of "Buffy Coat":
Depending on the type of tubes you have used (plastic ones are favorite, but also with glass tubes there is a possibility of breaking the glass by means of a glass cutting device and special pliers, which could be produced by a tool shop) you have to get rid of the bottom part of the tube. Remember that the bottom part (RBC's) will not have been fixed properly at that time (so use PSA, at least gloves!).
Cut the bottom part some mm below the RBC-layer, place tube into fixative again (swirling a little bit to get rid of loose and unfixed RBC's). then try to separate the whole layer-plug from the tube wall (there are some possibilities how to manage this) and try to push out the whole fixed plug.
Dissect under stereomicroscope: you certainly will see the different layers, among them the layer of thrombocytes / platelets right below the small fixed plasma layer.

Another possibility to collect platelets perhaps would be density gradient centrifugation (but this - with e.g. the SIGMA Histopaque®-Ficoll method usually is done to remove all platelets from the MNC-fraction).
Please also consider other techniques depending on the purpose of your intended studies.
If you have any question left, let me know
(also off list if you prefer... I have done a lot of those {buffy coats} in my "career"),
Best wishes and regards
Wolfgang

Wolfgang MUSS
EM-Lab
Pathology,
SALK-LKH (Gen. Hospital) SALZBURG
AUSTRIA




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} Gesendet: Mittwoch, 19. Dezember 2012 02:08
} An: Muß Wolfgang
} Betreff: [Microscopy] Prep for platelets for TEM
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} Email: jrkjack-at-aol.com
} Name: Judy King
} Organization: West Virginia University - Pathology
} Title-Subject: 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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From: tbargar-at-unmc.edu
Date: Tue, 29 Jan 2013 14:53:12 -0600
Subject: [Microscopy] flat embedding large size tissue in LR White

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear Listers;

I have a faculty member who wants to embed brain tissue pieces that are 100-200 microns thick, 2cm in length and 1 to 2cm in width in LR White. I have Aclar film and have located PTFE flat molds with large enough cavities that may work. Before I proceed I would like to hear from anybody who may have experience in doing this kind of embedding. Any advice is appreciated. I am especially interested in knowing if the Aclar film will really work well enough to seal the mold and prevent air from interfering with the polymerization. Also would it be best to first use UV polymerization in a cold chamber or can I go directly to the embedding oven at 65 degrees C? Could variable wattage microwave polymerization be used without submerging the mold underwater? As always thanks in advance for the help.

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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From: dsherman-at-purdue.edu
Date: Tue, 29 Jan 2013 15:03:21 -0600
Subject: [Microscopy] Re: flat embedding large size tissue in LR White

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Tom
Do you have access to an oven with vacuum? If so then just connect to a nitrogen source and alternate pulling a vacuum and the flooding with nitrogen to drive out the oxygen. Then leave the oven at 60oC to polymerize. Only leave vacuum low- at about 5lb - so as not to force resin to creep up the molds.
Debby

DS imaging LLC
Www.dsimagingllc.com

Sent from my iPhone

On Jan 29, 2013, at 3:55 PM, "tbargar-at-unmc.edu" {tbargar-at-unmc.edu} wrote:

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} Dear Listers;
}
} I have a faculty member who wants to embed brain tissue pieces that are 100-200 microns thick, 2cm in length and 1 to 2cm in width in LR White. I have Aclar film and have located PTFE flat molds with large enough cavities that may work. Before I proceed I would like to hear from anybody who may have experience in doing this kind of embedding. Any advice is appreciated. I am especially interested in knowing if the Aclar film will really work well enough to seal the mold and prevent air from interfering with the polymerization. Also would it be best to first use UV polymerization in a cold chamber or can I go directly to the embedding oven at 65 degrees C? Could variable wattage microwave polymerization be used without submerging the mold underwater? As always thanks in advance for the help.
}
} 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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From: duraine-at-bcm.edu
Date: Tue, 29 Jan 2013 16:16:28 -0600
Subject: [Microscopy] flat embedding large size tissue in LR White

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hello Tom,

Yes, LR White can be tricky, but we have a method that seems odd but always works. In our experience it is better to not fill all the PTFE mold spaces with samples. We have to orientate drosophila retina and larvae very strategically, so we place our samples into the center 6 wells, overfill the wells including two extra empty ones on either side of the samples. Press the Aclar film over the samples in the middle first then gently lay out the film toward both sides so that the resin pours over into the left-over empty well spaces toward the ends. The very end well spaces will not polymerise very well, but the ones with the samples will turn out very well, at least for us. We then place the mold into a 60 degree oven overnight to two days, sometimes three depending on the weather. So far we have been able to thick section and do fluorescence staining with no problem. Hope this helps.


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-----
X-from: tbargar-at-unmc.edu [mailto:tbargar-at-unmc.edu]
Sent: Tuesday, January 29, 2013 3:05 PM
To: Duraine, Lita

Dear Listers;

I have a faculty member who wants to embed brain tissue pieces that are 100-200 microns thick, 2cm in length and 1 to 2cm in width in LR White. I have Aclar film and have located PTFE flat molds with large enough cavities that may work. Before I proceed I would like to hear from anybody who may have experience in doing this kind of embedding. Any advice is appreciated. I am especially interested in knowing if the Aclar film will really work well enough to seal the mold and prevent air from interfering with the polymerization. Also would it be best to first use UV polymerization in a cold chamber or can I go directly to the embedding oven at 65 degrees C? Could variable wattage microwave polymerization be used without submerging the mold underwater? As always thanks in advance for the help.

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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18, 28 -- From duraine-at-bcm.edu Tue Jan 29 16:16:28 2013
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From: bigelow-at-umich.edu
Date: Tue, 29 Jan 2013 21:35:40 -0600
Subject: [Microscopy] Turbo pumps and oil contamination

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

One of the outstanding characteristics of turbomolecular pumps is
that they develop such high compression ratios for oil molecules that
these molecules are unable to move from the backing line, through the
pump, and into the vacuum system - provided the pump's rotor is
turning at nearly full rotational speed. This, combined with the fact
that they can produce pressures in the UHV range, makes them
particularly attractive as pumps on electron microscopes. HOWEVER, if
the turbo pump is backed by an oil-sealed rotary vane pump achieving
this contamination-free operation requires very careful adherence to
proper pump-down and venting procedures. This important matter is
discussed in considerable detail in Section 6.1.9b (pages 255 - 258)
of my book, "Vacuum Methods in Electron Microscopy".

Now-a-days, however, a more straightforward solution is to use an
oil-free backing pump, such as a scroll pump. If you replace a rotary
vane pump with a scroll pump it would be wise to also replace the
backing line with a new oil-free line.
--
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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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 30 Jan 2013 07:11:28 -0600
Subject: [Microscopy] viaWWW:RuO4 staining

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Email: trent-at-ornl.gov
Name: shawn reeves

Organization: Oak Ridge National Lab

Title-Subject: [Filtered] RuO4 staining

Message: Hi,

I will be setting up an area to stain polypropylene blocks with RuO4. I'll be using the kit sold
from SPI which consist of RuO2 hydrate and sodium periodate. I'm wondering if anyone could guide me
on the safety of this process and how to dispose of the product. i think this kit produces 0.67%
aqueous RuO4 and I will need the solution to be 0.5% to dispose of it. How should I treat
everything that has come in contact with the solution, vials, tissues etc. Also I will be slicing
the polypropylene with our microtome, is there any safety issues concerning the stained
polypropylene which will be falling in the microtome?

Thank you everyone in advance for your replies!

shawn

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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 30 Jan 2013 07:15:58 -0600
Subject: [Microscopy] viaWWW:Need manual for EM Tech K850 Critical point dryer

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Email: margaret.dienelt-at-ars.usda.gov
Name: Margaret Dienelt

Organization: Agricultural Research Service, USDA

Title-Subject: [Filtered] Need manual for EM Tech K850 Critical point dryer

Message: Hello Everyone,

We have an old EM Tech K850 critical point dryer in our lab that has never been used. After all
these years, we need to use it but of course the manual is nowhere to be found. Does anyone have a
manual to this instrument they can spare? If you have a manual but can't spare it, would you please
share the contact information of EM Tech with us?

Another option: if you can spare your manual for a week, we will copy it at our expense and send it
back the next day.

Thank you very much.

Margaret Dienelt
Plant Pathologist/Electron Microscopist
Floral and Nursery Plants Research Unit
NA/ARS/USDA

Bldg 012 Rm 1-7
10300 Baltimore Avenue
Beltsville, MD 20705 USA

Phone: 760-263-4219, 301-504-6097
Email: margaret.dienelt-at-ars.usda.gov
Fax: 301-504-5096



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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 30 Jan 2013 07:16:46 -0600
Subject: [Microscopy] viaWWW:Infiltrate EMbed 812 into gram positive bacteria

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Email: joseph.uknalis-at-ars.usda.gov
Name: Joe Uknalis

Organization: USDA-ERRC

Title-Subject: [Filtered] Infiltrate EMbed 812 into gram positive bacteria

Message: I've had difficulty getting good infiltration into Listeria and Staphlococcus. I've tried
long infiltration times (2 days) and under house vacuum.

Is the secret to good infiltration even longer times, higher vacuum levels or ???

thanks

Joe


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From: eschumacher-at-mccrone.com
Date: Wed, 30 Jan 2013 11:22:52 -0600
Subject: [Microscopy] Short Course Announcement: SEM

Contents Retrieved from Microscopy Listserver Archives
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Greetings Fellow Microscopists,

Hooke College of Applied Sciences, located in Westmont, IL, is offering a scanning electron microscopy short course March 4-8, 2013. 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)







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From: mraderma-at-uvm.edu
Date: Wed, 30 Jan 2013 15:32:55 -0600
Subject: [Microscopy] M&M meeting 2013, three interesting symposia

Contents Retrieved from Microscopy Listserver Archives
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Dear Colleagues,

We would like to bring to your attention three symposia at the
upcoming M&M 2013 meeting,
to be held in Indianapolis, Indiana on August 4-8, 2013. We would like
to encourage you
to submit a short paper and join us in Indianapolis.

This year the M&M conference will begin with a plenary session
featuring a talk by
Prof. Harald Rose, “The Long-Lasting Struggle to Achieve
Atomic-Resolution Microscopy
by Correcting the Aberrations of Electron Lenses". In his presentation
Prof. Rose will
provide an overview of the history of electron microscopy, and give
key insights into
the challenges of electron microscopy in the future. The second
keynote speaker will
be Joris Dik with a talk entitled: "Looking Through Paintings" With a
background in
Art History, Chemistry and Materials Sciences, Professor Dik brings a
unique perspective
to the study of paintings and masterworks, combining insights from
both the science
and the art worlds.

The three symposia related to structure, ultrastructure, and
technology development are:
_________________________________________________________________________________________
A04 Electron Tomography in Life and Material Science
Organizers: Heiner Friedrich, Montserrat Barcena, Esther Bullitt

Advances in sample preparation, instrumentation and methodology have
widened the scope
of electron tomography (ET) from sub-nanometer details to the
micrometer scale. This
symposium will address leading scientific and technological
developments in the physical
and biological sciences, using the widest possible range of ET imaging
approaches and
their integration with complementary techniques. Applications and
developments covering
— but not limited to — (aberration-corrected, energy-filtered) TEM
and STEM, phase plates,
diffraction or holography, are invited. Contributions including
complementary approaches
such as correlative light-electron microscopy, X-ray tomography,
serial-block face
SEM/AFM, and novel processing tools are encouraged.

Invited presenters: David Mastronarde, Elizabeth Wright, Irene Wacker,
Dirk van Dyck,
Krijn de Jong, Daniel Wolf
____________________________________________________________________________________________
A09 Advances in Data Processing in Optical and Electron Microscopy
Organizers: Edward P. Morris, David Morgan, Jeffrey L. Clendenon

Many advances in microscopy have been driven by developments in
software and the availability
of affordable high-speed computing. This symposium will focus on
software tools for EM and
LM that are publically available with an emphasis on analysis after
data acquisition. A major
topic will be EM software for structural biology (analysis of single
particles, helical
structures, 2d crystals), TEM/STEM tomography and enhancing
information obtained from
materials. Another focus will be software developed for LM to process,
visualize, segment
and measure 3D/4D images.

Invited presenters: Kevin Eliceiri, James Glazier, Michael
Radermacher, Hanspeter Winkler,
Quentin Ramasse, Lewys Jones

____________________________________________________________________________________________
B03 Structural Biology and Cell Ultrastructure
Organizers: Paula C. A. da Fonseca, Michael Radermacher, Ingeborg Schmidt-Krey

Our understanding of the 3D structure and function of cells,
microorganisms and macromolecular
assemblies has experienced great advances through recent developments
of EM techniques and
hybrid methodologies. This symposium highlights structural and
ultrastructural studies of cells,
microorganisms and biological macromolecules using electron microscopy
techniques
(e.g. single-particle analysis, tomographic methods; helical
reconstruction, crystallographic
methods) singly or combined with other structural methods (e.g. X-ray
methods; atomic force
microscopy). Topics will include: structure and function of
macromolecular assemblies, virus
structure and virus-host interactions; eukaryotic and prokaryotic cell
architecture; cellular
metabolism; cell division and protein translation; cellular secretion,
adhesion and motility;
cell-cell communication and signaling.

Invited presenters: David Veesler, Edward Morris, Beate Rockel, Yifan
Cheng, Tamir Gonen,
Timothy Baker
_____________________________________________________________________________________________
Please also spend some time to browse the complete program and you
will discover many other
presentation of high interest to our field.

Specifics:
The abstract/short paper submission deadline is on February 15th. The
submission website will
close at 5PM PST on this date. Contributions must be 2-pages long and
have at least 300 words.
Note that 1-page or shorter contributions might be rejected and, if
accepted for presentation,
will not be part of the proceedings. The 2-page meeting contributions
can be deposited under
the public access policy of granting agencies (e.g. Pubmedcentral / NIH).

When you submit your paper, select any of the three symposia:
A04 Electron Tomography in Life and Material Science
A09 Advances in Data Processing in Optical and Electron Microscopy
B03 Structural Biology and Cell Ultrastructure

Please see the full paper submission guidelines on the meeting website.
Link to meeting site: http://www.microscopy.org/MandM/2013/callforpapers.cfm
This is a great educational and networking meeting for the entire
microscopy community.
Students can benefit from a reduced registration fee ($50.00) and have
ample opportunities
to receive awards either before the meeting (make sure to check the
corresponding box during
submission) or during the poster session. For more information go to:
http://www.microscopy.org/MandM/2013/callforpapers.cfm

When registering, please also check out the many interesting Sunday
pre-meeting courses
and workshops.

We look forward to seeing many of you at the meeting.


Sincerely,
The organizers of A04, A09, and B04


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From: beth-at-plantbio.uga.edu
Date: Wed, 30 Jan 2013 15:34:52 -0600
Subject: [Microscopy] cpds and sputter-coaters

Contents Retrieved from Microscopy Listserver Archives
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Hi all,
I would like your advice/opinion on critical point dryers and sputter-
coaters. We are leaning towards the Tousimis 931 cpd (winner of the
2012 Microscopy Today Innovation Award) and the SPI Module sputter/
carbon-coaters.
thanks in advance,
Beth


==============================Original Headers==============================
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From: pekysar-at-ucdavis.edu
Date: Wed, 30 Jan 2013 16:11:15 -0600
Subject: [Microscopy] cpds and sputter-coaters

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi Beth,
We recently purchased the Tousimis 931. I absolutely love it!!!!!!! It's
awesome. The samples I processed have been superb!!! It has tremendously
freed up my time and has been completely dependable.
I'm not familiar with the SPI sputter/carbon coaters. We have a PELCO SC-7
Sputter coater and it's probably the best one I've ever used. It's easy and
quick. However it is just a low vacuum sputter coater(0.08mbar).
Hope that helps!
Pat Kysar, C.E.M.T.
Chair, Certification Board, Microscopy Society of America
University of California, Davis
School of Medicine
Dept of Pathology and Laboratory Medicine
Electron Microscopy Lab



-----Original Message-----
X-from: beth-at-plantbio.uga.edu [mailto:beth-at-plantbio.uga.edu]
Sent: Wednesday, January 30, 2013 1:48 PM
To: pekysar-at-ucdavis.edu

Hi all,
I would like your advice/opinion on critical point dryers and sputter-
coaters. We are leaning towards the Tousimis 931 cpd (winner of the
2012 Microscopy Today Innovation Award) and the SPI Module sputter/
carbon-coaters.
thanks in advance,
Beth


==============================Original Headers==============================
2, 20 -- From beth-at-plantbio.uga.edu Wed Jan 30 15:34:52 2013
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12, 30 -- From pekysar-at-ucdavis.edu Wed Jan 30 16:11:15 2013
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From: Rosemary.White-at-csiro.au
Date: Wed, 30 Jan 2013 16:33:29 -0600
Subject: [Microscopy] cpds and sputter-coaters

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

We'd have to second that, we got the earlier model Tousimis 815 and it has been very good though rather greedy for CO2. It would be worth looking at the new fully automatic Leica CPD, looked good on demo. Fully automatic is the only way...

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
________________________________________
X-from: pekysar-at-ucdavis.edu [pekysar-at-ucdavis.edu]
Sent: Thursday, 31 January 2013 9:14 a.m.
To: White, Rosemary (PI, Black Mountain)

Hi Beth,
We recently purchased the Tousimis 931. I absolutely love it!!!!!!! It's
awesome. The samples I processed have been superb!!! It has tremendously
freed up my time and has been completely dependable.
I'm not familiar with the SPI sputter/carbon coaters. We have a PELCO SC-7
Sputter coater and it's probably the best one I've ever used. It's easy and
quick. However it is just a low vacuum sputter coater(0.08mbar).
Hope that helps!
Pat Kysar, C.E.M.T.
Chair, Certification Board, Microscopy Society of America
University of California, Davis
School of Medicine
Dept of Pathology and Laboratory Medicine
Electron Microscopy Lab



-----Original Message-----
X-from: beth-at-plantbio.uga.edu [mailto:beth-at-plantbio.uga.edu]
Sent: Wednesday, January 30, 2013 1:48 PM
To: pekysar-at-ucdavis.edu

Hi all,
I would like your advice/opinion on critical point dryers and sputter-
coaters. We are leaning towards the Tousimis 931 cpd (winner of the
2012 Microscopy Today Innovation Award) and the SPI Module sputter/
carbon-coaters.
thanks in advance,
Beth


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From: wa5ekh-at-juno.com
Date: Wed, 30 Jan 2013 18:59:44 -0600
Subject: [Microscopy] Digital Camera adapter for Slow Scan Photography-Polaroid SEM Fixture

Contents Retrieved from Microscopy Listserver Archives
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I need some details on attaching and operating a Digital camera to the Slow Scan Photography-Polaroid Fixture on an old SEM. The college doesn't seem to have enough funding for the commercial camera fixtures.
Jeff/N. Texas Area
____________________________________________________________
Woman is 57 But Looks 27
57-Year-Old Mom has a simple facelift trick that angered doctors...
http://thirdpartyoffers.juno.com/TGL3131/5109c1f678c8241f619bbst02vuc


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From: stefan.diller-at-t-online.de
Date: Thu, 31 Jan 2013 03:55:18 -0600
Subject: [Microscopy] NORAN SIX jumper-settings

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear All,
anybody out there who can help me with the correct jumper settings for the scan voltage at the NORAN SIX EDS system?
I need to have ca 8 Volt Vss for my scan-amp, now I have only 5 Vss. With the potis at the rear side I am not able to go up high
enough...

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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From: tbargar-at-unmc.edu
Date: Thu, 31 Jan 2013 10:40:23 -0600
Subject: [Microscopy] Thick sectioning the large LR White embedded tissue

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear Listers:

My thanks to everyone who responded to my request on flat embedding a large piece of tissue (2cm long, 1 to 1.5cm wide and 100microns thick) in LR White. So the embedding certainly looks feasible. Related to this project, the faculty member has asked me if it is possible to section the resulting block in 10 micron thick sections? This is for confocal imaging apparently. So he would like to get a section that is 10microns thick and approx. 2cm long and 1 to 1.5cm wide and I presume heat fixed to a glass slide. Now the thickest I have ever cut is 2 microns on my Diatome Jumbo Histo diamond and less than 8mm wide. Anyone out there with experience in cutting a section like this? I feel the LR White would be too brittle to go as thick as 10 microns without cracking, but I really don't know. Can the block be sectioned on a standard histology microtome, the way you section a paraffin block? Any and all advice would be appreciated.

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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From: dsherman-at-purdue.edu
Date: Thu, 31 Jan 2013 11:00:01 -0600
Subject: [Microscopy] Re: Thick sectioning the large LR White embedded tissue

Contents Retrieved from Microscopy Listserver Archives
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Tom,

Do you have the option of embedding some of the tissue in another resin
that would not be suitable for TEM? If so than I would suggest embedding
in JB-4 resin which can be cut with larger glass knives (and also
disposable metal blades). Usually this is done on a JB-4 microtome and
uses wider glass that is cut with a special knife breaker. These pieces
of equipment may not be available to you. However, it may be possible to
use a standard paraffin microtome to cut sections of this size in this
resin. Perhaps others can comment on that since I have not tried it.

Ultrastructure of tissue embedded in JB-4 resin is of much higher quality
than that embedded in paraffin. It also can still be stained with many of
the aqueous stains used for LM.

Debby


Debra Sherman, Founder & CSO
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 1/31/13 11:42 AM, "tbargar-at-unmc.edu" {tbargar-at-unmc.edu} wrote:

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From: parishcm-at-ornl.gov
Date: Thu, 31 Jan 2013 13:59:29 -0600
Subject: [Microscopy] Postdoc position open at Oak Ridge National Laboratory

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Sent on behalf of Dr. M. K. Miller, address for queries below.

------

A new postdoc position is available immediately at Oak Ridge National Laboratory

Purpose
Under general supervision, the incumbent will focus on the nanostructural characterization of the microstructures of
nanostructured ferritic alloys under various high dose ion irradiations and thermally aged treatments. This position resides in
the Microscopy Group in the Materials Science and Technology Division (MSTD), Physical Sciences Directorate (PSD) at
Oak Ridge National Laboratory.

Major Duties/Responsibilities
- Perform atom probe tomography, scanning electron microscopy, and transmission electron microscopy
- Responsible for the operation of the local electrode atom probe and transmission electron microscopes
- Prepares specimens with a dual-beam scanning electron microscope and focused ion beam miller
- Performs atom probe tomography data reconstruction and analysis
- Ensures compliance with environment, safety, health and quality program requirements.
- Maintains strong commitment to the implementation and perpetuation of values and ethics.

(more details are in the attachment)

To apply:
Go to http://jobs.ornl.gov/
Select View External Positions button on the rightmost column
Enter "tomography" in the keywords and hit start button
Double clink on Postdoctoral Research Associate / NB50344514

See the buttons at the top of the page to apply.

For addition details, please e-mail (millermk-at-ornl.gov)
Dr. Michael K Miller
Corporate Fellow

Microscopy Group
Materials Science and Technology Division,
Oak Ridge National Laboratory
P. O. Box 2008 (please replace POB with 1 Bethel Valley Road for express mail)
Oak Ridge, TN 37831-6139, USA

Phone: (865) 574 4719
Fax: (865) 241 3650


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From: larry.ackerman-at-ucsf.edu
Date: Thu, 31 Jan 2013 14:24:08 -0600
Subject: [Microscopy] Re: Thick sectioning the large LR White embedded

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Hi Tom,
It doesn't make sense to me to epoxy embed and cut 10um sections for
confocal examination. For confocal I suggest cutting the large tissue at
50 um then process and image with a confocal. Most confocals can image
at a depth of at least 50um. A multi-photon confocal can image deeper
~200-300um. Otherwise if the epoxy preparation is important cut 2um
sections and use a widefield microsccope--confocal is not really needed
since you already have thin sections. The individual section images can
be aligned post-microscopy. However, considering the problems of section
folding and wrinkling that I have encountered with 4mm X 4mm X 1.5mm
epoxy sections, larger sections may present even more headaches and
frustration.
Good luck,
Larry

On 1/31/2013 8:51 AM, tbargar-at-unmc.edu wrote:
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} Dear Listers:
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} My thanks to everyone who responded to my request on flat embedding a large piece of tissue (2cm long, 1 to 1.5cm wide and 100microns thick) in LR White. So the embedding certainly looks feasible. Related to this project, the faculty member has asked me if it is possible to section the resulting block in 10 micron thick sections? This is for confocal imaging apparently. So he would like to get a section that is 10microns thick and approx. 2cm long and 1 to 1.5cm wide and I presume heat fixed to a glass slide. Now the thickest I have ever cut is 2 microns on my Diatome Jumbo Histo diamond and less than 8mm wide. Anyone out there with experience in cutting a section like this? I feel the LR White would be too brittle to go as thick as 10 microns without cracking, but I really don't know. Can the block be sectioned on a standard histology microtome, the way you section a paraffin block? Any and all advice would be appreciated.
}
} 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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--
Larry Ackerman, Specialist
Electron Microscopy Lab
Manager, Diabetes & Endocrinology Research Center Microscopy Core
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From: mary.raven-at-lifesci.ucsb.edu
Date: Thu, 31 Jan 2013 16:27:13 -0600
Subject: [Microscopy] convention microscope service and repair providers

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Dear List
I frequently receive requests for recommendations of reliable service
providers for conventional microscopes. Issues like cleaning crusted
optics, repairing and replacing springs etc. Are there any you can
recommend that work in Southern California, USA?

Thanks
Mary

--
Mary Raven
Microscopy Facility Director
http://www.lifesci.ucsb.edu/~m_raven/
Phone: (805) 893 8702

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

2014 Advanced Microscopy and Digital Imaging Workshop
https://www.lifesci.ucsb.edu/mcdb/events/imaging_workshop/
Jan 14- Jan 17, 2014



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From: mike-at-boeckeler.com
Date: Fri, 1 Feb 2013 14:47:31 -0600
Subject: [Microscopy] Work Shop

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Hello,
We would like to invite you to an ultra microtomy work shop hosted by
San Joaquin Delta College, Stockton, on February 25th-28th. We will
cover both room temp and cryo processes for material and biological
specimen sample prep. There will be lectures in the morning followed by
"hands-on" training after lunch. We encourage our attendees to bring a
sample of what they are currently working on. SJDC - Stockton has a well
equipped imaging facility and RMC will have the latest cryo
ultramicrotome and glass knife maker to use. There is no charge to
attend the workshop, lunch is included, but attendees will be
responsible for their own travel and accomodation expenses. Space is
limited so please let me know as soon as possible. February 7th is the
sigh up deadline.
Thank you,
Mike Leeper

--
Mike Leeper
RMC National Sales and Service Manager
Boeckeler Instruments, Inc.
4650 S. Butterfield Drive
Tucson, AZ U.S.A. 85714
Phone: (520) 745-0001 Fax: (520) 745-0004

mike-at-boeckeler.com
www.rmcproducts.com


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From: WHITTAKS-at-si.edu
Date: Sat, 2 Feb 2013 08:33:39 -0600
Subject: [Microscopy] viaWWW:Need manual for EM Tech K850 Critical point dryer

Contents Retrieved from Microscopy Listserver Archives
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Email: margaret.dienelt-at-ars.usda.gov
Name: Margaret Dienelt

Organization: Agricultural Research Service, USDA

Title-Subject: [Filtered] Need manual for EM Tech K850 Critical point dryer

Message: Hello Everyone,

We have an old EM Tech K850 critical point dryer in our lab that has never been used. After all these years, we need to use it but of course the manual is nowhere to be found. Does anyone have a manual to this instrument they can spare? If you have a manual but can't spare it, would you please share the contact information of EM Tech with us?

Another option: if you can spare your manual for a week, we will copy it at our expense and send it back the next day.

Thank you very much.

Margaret Dienelt
Plant Pathologist/Electron Microscopist
Floral and Nursery Plants Research Unit
NA/ARS/USDA

Bldg 012 Rm 1-7
10300 Baltimore Avenue
Beltsville, MD 20705 USA

Phone: 760-263-4219, 301-504-6097
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From: larry.ackerman-at-ucsf.edu
Date: Mon, 4 Feb 2013 14:12:18 -0600
Subject: [Microscopy] Thick sectioning the large LR White

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I haven't used Spurr's for many years. I use Eponate 812 or EMbed--Epon
equivalents. I put a drop of water on a glass slide then transfer a
section from the knife boat to the drop of water with a Minutien pin
(very fine needle) glued on a wood applicator stick. Usually I put two
or three sections on a drop but for larger sections one per drop. The
slide then goes on a hot plate set at a temperature so the water drop
dries slowly ~3-4 minutes. Sometimes just allowing the water to air dry
at room temperature works better. Then I stain the sections with
Toluidine Blue for 10-20 seconds and rinse with water. I am usually
just looking for an area that I wish to thin section and do not use the
1-2um sections for microscopy. Another possibility to minimize wrinkles
is to stretch the sections in the knife boat--with a heat pen or solvent
vapors (chloroform vapors are not healthy).
Happy Sectioning!
Larry

On 1/31/2013 12:52 PM, Philip Oshel wrote:
} Larry,
}
} How do you deal with section wrinkling & folding in large sections?
} I've got a student do LM on corn kernels - embryonic development - and
} he's reached a stage where he needs large area sections. Not 4x4 mm,
} but a couple of millimeters, maybe more. Using Spurr's, I believe.
} That's what I have, and I suspect the prof he's working with is also
} using that, not some acrylate or the like.
} This is for LM and tol-blue staining, although they're moving towards
} doing some TEM.
}
} Phil
}
} }
} } Hi Tom,
} } It doesn't make sense to me to epoxy embed and cut 10um sections for
} } confocal examination. For confocal I suggest cutting the large tissue at
} } 50 um then process and image with a confocal. Most confocals can image
} } at a depth of at least 50um. A multi-photon confocal can image deeper
} } ~200-300um. Otherwise if the epoxy preparation is important cut 2um
} } sections and use a widefield microsccope--confocal is not really needed
} } since you already have thin sections. The individual section images can
} } be aligned post-microscopy. However, considering the problems of section
} } folding and wrinkling that I have encountered with 4mm X 4mm X 1.5mm
} } epoxy sections, larger sections may present even more headaches and
} } frustration.
} } Good luck,
} } 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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From: henning.stahlberg-at-unibas.ch
Date: Wed, 6 Feb 2013 03:16:21 -0600
Subject: [Microscopy] 3DEM GRC 2014 will be June 22-27 at Melia Golf Vichy Catalan,

Contents Retrieved from Microscopy Listserver Archives
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Dear Colleagues,

The 3DEM GRC in 2014 will take place from June 22 to 27, 2014, in Girona, Spain. This is close to Barcelona.

Please mark your calendars.

Henning.

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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From: Pete.Finger-at-jax.org
Date: Thu, 7 Feb 2013 14:05:17 -0600
Subject: [Microscopy] Facility Information

Contents Retrieved from Microscopy Listserver Archives
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Scientific Services at The Jackson Laboratory is conducting benchmarking surveys in an effort to develop an understanding of the current best practices in operations, management and technical delivery at peer institutions. The goal is to gather quality data that can be used to assist us and all survey participants with operational assessment and planning. Survey participants will receive compiled results which will be de-identified before distribution. Thank you in advance for your participation.


Pete Finger
Manager, Electron Microscopy Service
The Jackson Laboratory
Bar Harbor, ME 04609
207-288-6337

"Do your duty in all things. You can not do more, you should never wish to do less".
- Robert E. Lee




The information in this email, including attachments, may be confidential and is intended solely for the addressee(s). If you believe you received this email by mistake, please notify the sender by return email as soon as possible.


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From: eric-miller-at-northwestern.edu
Date: Fri, 8 Feb 2013 14:20:51 -0600
Subject: [Microscopy] New Series: What's It Made Of?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

The NUANCE Center at Northwestern University is going to be producing a new web series called "What's It Made Of?" It will be a short, semi-educational show mainly featuring SEM and EDS to examine normal, everyday objects. We'd love to use this as a fun educational outreach to show the uninitiated the types of things that we can do in the world of electron microscopy. So please share with all your friends and whoever.

The first episode has been posted and examines the sparkly ink on the $20 bill.

http://youtu.be/z3qHg1QOAk4


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




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From: tindallr-at-missouri.edu
Date: Fri, 8 Feb 2013 16:44:43 -0600
Subject: [Microscopy] Gatan manuals?

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear Collective,

Might someone out there have manuals for a Gatan 601 Ultrasonic Disc Cutter=
and Gatan 656 Dimple Grinder that they would be willing to share, copy, le=
nd, or PDF? We have two lonely instruments waiting for some attention, but=
we just don't know their proper care and feeding, yet. Any assistance wou=
ld be greatly appreciated and postage, copying, etc. costs reimbursed with =
all dispatch.

Thanks!

Randy
(from home----signature's on the office computer----too lazy to add one her=
e now)

==============================Original Headers==============================
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From: tindallr-at-missouri.edu
Date: Sat, 9 Feb 2013 10:34:49 -0600
Subject: [Microscopy] Gatan manuals!

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Thanks to all who responded to our request for manuals. We have either received the ones we need, or they are on their way. What a great bunch you guys are!

Randy

==============================Original Headers==============================
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From: tindallr-at-missouri.edu
Date: Mon, 11 Feb 2013 10:10:27 -0600
Subject: [Microscopy] Cryo-ultramicrotomy: trimming hpf cups

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear Collective,

I know. I'm getting to be a real pest, but the day of reckoning has come. We need to take sequential sections down through an HPF-frozen sample and doubt that we will be able to get the sample out of the cup. This means we will probably have to trim away the cup from around the sample. We know this can be done (in fact, we are trimming away gold with an old beat-up diamond as I write this), but doing it in a time frame that makes it affordable and doesn't destroy the knife in the process is the problem.

A couple potential approaches:

1) pretrimming the block with a dremel or similar tool and leaving a small amount of gold intact around the sample cup area before doing HPF. Would this still provide an adequate seal during the actual freezing?

2) doing the trimming of the metal in a cryostat with a steel knife after freezing the sample, then transferring into the cryo-ultramicrotome for the final sections.

Does anyone have any thoughts on these or other approaches? To repeat, we hope to get the cup-trimming part down to a manageable amount of time, without trashing every (expensive) knife we have.

Thanks!

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)



==============================Original Headers==============================
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13, 30 -- "Vinson, Juliana Pavlovna"
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From: PhillipsT-at-missouri.edu
Date: Mon, 11 Feb 2013 10:35:59 -0600
Subject: [Microscopy] Cryo-ultramicrotomy: trimming hpf cups

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I doubt a cryostat would be cold enough to prevent crystallization of the of vitrified ice.

If a diamond can cut the gold, have you tried a glass knife?

Good luck and please share the final method that leads to success. 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: tindallr-at-missouri.edu [mailto:tindallr-at-missouri.edu]
Sent: Monday, February 11, 2013 10:11 AM
To: Phillips, Thomas E.

Dear Collective,

I know. I'm getting to be a real pest, but the day of reckoning has come. We need to take sequential sections down through an HPF-frozen sample and doubt that we will be able to get the sample out of the cup. This means we will probably have to trim away the cup from around the sample. We know this can be done (in fact, we are trimming away gold with an old beat-up diamond as I write this), but doing it in a time frame that makes it affordable and doesn't destroy the knife in the process is the problem.

A couple potential approaches:

1) pretrimming the block with a dremel or similar tool and leaving a small amount of gold intact around the sample cup area before doing HPF. Would this still provide an adequate seal during the actual freezing?

2) doing the trimming of the metal in a cryostat with a steel knife after freezing the sample, then transferring into the cryo-ultramicrotome for the final sections.

Does anyone have any thoughts on these or other approaches? To repeat, we hope to get the cup-trimming part down to a manageable amount of time, without trashing every (expensive) knife we have.

Thanks!

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)



==============================Original Headers==============================
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From: PhillipsT-at-missouri.edu
Date: Mon, 11 Feb 2013 12:46:01 -0600
Subject: [Microscopy] Cryo-ultramicrotomy: trimming hpf cups

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I just remembered - I think the cryo-ultramicrotome has a tool for this! It is a metal rod with a sharp end that fits in the knife holder - take a peek in your goodies box. you could have extras made at the machine shop


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 doubt a cryostat would be cold enough to prevent crystallization of the of vitrified ice.

If a diamond can cut the gold, have you tried a glass knife?

Good luck and please share the final method that leads to success. 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: tindallr-at-missouri.edu [mailto:tindallr-at-missouri.edu]
Sent: Monday, February 11, 2013 10:11 AM
To: Phillips, Thomas E.

Dear Collective,

I know. I'm getting to be a real pest, but the day of reckoning has come. We need to take sequential sections down through an HPF-frozen sample and doubt that we will be able to get the sample out of the cup. This means we will probably have to trim away the cup from around the sample. We know this can be done (in fact, we are trimming away gold with an old beat-up diamond as I write this), but doing it in a time frame that makes it affordable and doesn't destroy the knife in the process is the problem.

A couple potential approaches:

1) pretrimming the block with a dremel or similar tool and leaving a small amount of gold intact around the sample cup area before doing HPF. Would this still provide an adequate seal during the actual freezing?

2) doing the trimming of the metal in a cryostat with a steel knife after freezing the sample, then transferring into the cryo-ultramicrotome for the final sections.

Does anyone have any thoughts on these or other approaches? To repeat, we hope to get the cup-trimming part down to a manageable amount of time, without trashing every (expensive) knife we have.

Thanks!

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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From: bbandli-at-d.umn.edu
Date: Mon, 11 Feb 2013 14:36:43 -0600
Subject: [Microscopy] JEOL aperture problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

As Tom Phillips suggests, there are options in the cryoultramicrotome. We do trimming of frozen roots with attached soil with a tungsten-coated glass knife, which would probably work for you.

Also, could you make/buy the freezing cups in a softer metal or other material?

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
________________________________________
X-from: tindallr-at-missouri.edu [tindallr-at-missouri.edu]
Sent: Tuesday, 12 February 2013 3:17 a.m.
To: White, Rosemary (PI, Black Mountain)

Dear Collective,

I know. I'm getting to be a real pest, but the day of reckoning has come. We need to take sequential sections down through an HPF-frozen sample and doubt that we will be able to get the sample out of the cup. This means we will probably have to trim away the cup from around the sample. We know this can be done (in fact, we are trimming away gold with an old beat-up diamond as I write this), but doing it in a time frame that makes it affordable and doesn't destroy the knife in the process is the problem.

A couple potential approaches:

1) pretrimming the block with a dremel or similar tool and leaving a small amount of gold intact around the sample cup area before doing HPF. Would this still provide an adequate seal during the actual freezing?

2) doing the trimming of the metal in a cryostat with a steel knife after freezing the sample, then transferring into the cryo-ultramicrotome for the final sections.

Does anyone have any thoughts on these or other approaches? To repeat, we hope to get the cup-trimming part down to a manageable amount of time, without trashing every (expensive) knife we have.

Thanks!

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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From noreply-at-creatividentro.com Mon Feb 11 13:54:07 2013
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Hello,

I am having an issue with the moveable aperture (part MP-30060) on my
JSM-6490LV and am hoping someone out there has had a similar issue and can
be kind enough to provide some insight.

The mechanism has coarse positioning mechanism to hold the objective
aperture strip in three positions plus a fully extracted position.
Currently, it is only able to be positioned in the middle two positions
(aperture positions #2, and #1). Any help would be appreciated.

Thanks in advance,
Bryan


--
_________________________________________________________________
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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7, 44 -- Subject: JEOL aperture problem
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From: wesaia-at-iastate.edu
Date: Mon, 11 Feb 2013 15:52:26 -0600
Subject: [Microscopy] JEOL aperture problem

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

My experience with JEOL (and many other) aperture carriers is that the fine adjustment is enough to (nearly) shift from one aperture to another. So instead of seeing apertures 1, 2, 3, and OPEN you end up seeing nothing, 1, 2, and OPEN or 2, 3, nothing, and OPEN. If you have an ammeter on your system or an EDS system, you can check eventually determine which situation you have. You would have to know what the normal current or x-ray signal is for a given aperture and condenser lens setting.

I would experiment, carefully, with the longitudinal adjustment to see of you can bring the next aperture into view. You should then find all three apertures in their usual places.

BTW, I had trouble with a user damaging our strip. I tried to adjust my x-ray count rate but there was no difference between the positions. It turned out that they forgot the order of the twist and pull operations, and they put a 45-degree twist in the aperture strip. They dropped the strip down into the column. At lease it did not block the beam. Your problem is considerable simpler

Warren.
________________________________________
X-from: bbandli-at-d.umn.edu [bbandli-at-d.umn.edu]
Sent: Monday, February 11, 2013 2:37 PM
To: wesaia-at-iastate.edu

Hello,

I am having an issue with the moveable aperture (part MP-30060) on my
JSM-6490LV and am hoping someone out there has had a similar issue and can
be kind enough to provide some insight.

The mechanism has coarse positioning mechanism to hold the objective
aperture strip in three positions plus a fully extracted position.
Currently, it is only able to be positioned in the middle two positions
(aperture positions #2, and #1). Any help would be appreciated.

Thanks in advance,
Bryan


--
_________________________________________________________________
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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14, 39 -- Subject: RE: [Microscopy] JEOL aperture problem
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From: CrockerV-at-ninds.nih.gov
Date: Tue, 12 Feb 2013 13:37:09 -0600
Subject: [Microscopy] Soft epoxy blocks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Good Afternoon,

Does anyone have any tips for hardening some epoxy blocks that are too soft to section? Somehow, they didn’t harden as well as they usually do. I’ve put them back in the oven for a longer period of time, and they did get a bit harder, but not enough. They “bend”!! This has happened to me only twice in over 23 years! I’d really like to be able to section the samples (Fly embryos) for the PI.

Thanks,
Virginia Crocker
Biologist, NIH EM Facility



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From: ALawrence-at-i2at.msstate.edu
Date: Tue, 12 Feb 2013 16:43:04 -0600
Subject: [Microscopy] M&M 2013 meeting student bursary program

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

As the deadline for abstract submission approaches and plans are being
made to attend the Indianapolis meetings Aug. 4-8, please don*t forget
about MSA*s student bursary program. Its purpose is to encourage
students to attend the meetings by helping to defray some of the costs
and giving them an opportunity to meet and interact with the established
microscopy community.

The students will be paid $10 an hour to work for ~20 hours (up to 40
hours) during the meeting or pre-meeting events. 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 Café, and
helping with vendor tutorial sign-up. Payment is given as a check at
the end of the meetings or when the student leaves Indianapolis.

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 to help with meals
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 in section 2
of 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 their
registration fee.

For those *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 a meeting shirt
and the same cash allotment to help with meals. They also 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
662-325-7998
alawrence-at-i2at.msstate.edu





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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:10:17 -0600
Subject: [Microscopy] viaWWW:Student Petrographic Microscope

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Email: kenhon-at-hawaii.edu
Name: Ken Hon

Organization: University of Hawaii at Hilo

Title-Subject: [Filtered] Student Petrographic Microscope

Message: Aloha,

We are thinking about slowly replacing our aging Leitz monocular student
petrographic microscopes used in mineralogy and petrology. I was
wondering if anyone had suggestions for relatively inexpensive, but
durable binocular petrographic microscopes for use by students. Also,
they must withstand relatively high humidity. Even in a cabinet with a
dehumidifier stick in an air conditioned lab, the lowest we can get
humidity is about 50%. The Leitz's have lasted over 20 years and are
still workable, so they've been great from that standpoint. However,
they no longer make parts for them and we can't get them serviced any
longer. So, I'd welcome any ideas and suggestions for dealers (if
that's allowed on this board).

Aloha,

Ken



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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:11:36 -0600
Subject: [Microscopy] viaWWW:DIRECTOR, INTEGRATED MICROSCOPY & GRAPHICS EXPERTISE (IMAGE)

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Email: IMAGE-at-siu.edu
Name: George Trifon

Organization: Southern Illinois University

Title-Subject: [Filtered] DIRECTOR, INTEGRATED MICROSCOPY & GRAPHICS
EXPERTISE (IMAGE) FACILITY

Message: DIRECTOR, INTEGRATED MICROSCOPY & GRAPHICS EXPERTISE (IMAGE)
FACILITY
(Deadline Extended - 12 mo. Term, A/P)

Southern Illinois University Carbondale seeks to hire a Director for the
Integrated Microscopy & Graphic Expertise Facility. The Director will be
responsible for operating and maintaining state-of-the-art microscopy
equipment, teaching SEM and TEM classes, printing large posters for
research presentations, generating external contracts and grants,
overseeing billing and inventory control, and engaging in outreach and
promotional activities across campus and in the community.

Minimum Qualifications: Applicants must have a MasterÂ’s degree in
Science or Engineering and four (4) years experience as a professional
electron microscopist in a centralized, service-oriented facility with
multidisciplinary clients.

Preferred Qualifications: Experience in trouble-shooting with advanced
instrumentation. A working knowledge of graphic programs for use in
professional publications and large format posters. The ability to work
independently with minimal administrative oversight or support.
Excellent organizational and communication skills. Ph.D. preferred.

Website: image.siu.edu

Extended Deadline: Application review will begin Feb 18, 2013, and
continue until filled

Application: Send letter of application, resume, and three reference

letters via the website or hard copy to:

Chair, Search Committee
Office of Sponsored Projects Administration
SIU Carbondale
900 S Normal, Woody Hall C206, MC 4709
Carbondale, IL 62901

SIU Carbondale is an affirmative action/equal opportunity employer that
strives to enhance its ability to develop a diverse faculty and staff
and to increase its potential to serve a diverse student population. All
applications are welcomed and encouraged and will receive consideration.

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:12:39 -0600
Subject: [Microscopy] viaWWW:Electron microscopy schools?

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Email: serene_ng-at-dsi.a-star.edu.sg
Name: Serene

Title-Subject: [Filtered] Electron microscopy school

Message: Dear Listers,

I am looking for electron microscopy school(s) which offer short or
full-term courses (TEM in particular) - both theory and practical
lessons will be good.

I know there's Lehigh and Hooke which offer short courses. Are there
other institutions fellow listers can suggest? Are there similar schools
hosted in Europe or
Asia?

Thank you listers!
Serene

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:14:02 -0600
Subject: [Microscopy] viaWWW:CCD camera sensor size

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

Organization: IISc

Title-Subject: [Filtered] CCD camera sensor size

Message: Good evening,
I was wondering if anyone knows any way to find out exact sensor size of
ccd camera? (i want to measure actual distance, like in films, between
diffraction spots, any other suggestions are also welcome!)
i read the brochure of manufacturer, many things dont make sense there,
e.g chip size- 2/3 inches, 10.2mmx8.3mm
but wont 10.3mm x 8.3 mm be 1.31 cm diagonally? while 2/3 inches should
be 1.69 cm.
pixcel count-1376x 1032

thank you

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:14:58 -0600
Subject: [Microscopy] viaWWW:microtome accuracy

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Email: trogadisj-at-smh.ca
Name: Judy Trogadis

Organization: St. Michael's Hospital

Title-Subject: [Filtered] microtome accuracy

Message: Fellow microscopists

We are planning to do very fine optical measurements from tissue
sections, therefore, control of thickness and of even surface is critical.

Which are the most accurate and reproducible microtomes on the market
for paraffin or frozen blocks? Are there laser based tissue sectioning
equipment.

Any suggestions are welcomed, including replies from manufacturers.
Thank you,
Judy


Judy Trogadis
Bio-Imaging Coordinator
Keenan Research Centre, St. Michael's
209 Victoria Street
Toronto M5B 1T8, Canada
office: 416-864-6060 ext. 77612
imaging facility: ext. 77434
cell: 416-254-9330
trogadisj-at-smh.ca


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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:15:44 -0600
Subject: [Microscopy] viaWWW:LKB Manual needed

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Email: yankova-at-neuron.uchc.edu
Name: Maya Yankova

Organization: University of Connecticut Health Center

Title-Subject: [Filtered] LKB Manual needed

Message: Hello,
A researcher in our lab inherited an LKB microtome model 8800, but the
manual is missing. We were wandering if anyone has a pdf or hard copy
they could send us.

Thank you,

Maya Yankova
University of Connecticut,Health center

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:16:35 -0600
Subject: [Microscopy] viaWWW:Refurbished DoubleTilt Holder

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Title-Subject: [Filtered] Refurbished DoubleTilt Holder

Message: Is there any company that sells refurbished or used TEM holders?

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:17:21 -0600
Subject: [Microscopy] viaWWW:Fixation and Processing of Mouse Dorsal Root Ganglia

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Email: karen.rothberg-at-utsouthwestern.edu
Name: Karen Rothberg

Organization: UT Southwestern Medical Center

Title-Subject: [Filtered] Fixation and Processing of Mouse Dorsal Root
Ganglia

Message: Hi,

We have a user who wants 0.5 micron cross sections of the L3 ganglia
and spinal chords from mice. What is the best way to process the tissue
without mechanical damage to the root?

They did perfuse the mouse with Paraformaldehye/Glut and gave the
samples to us in 2.5% glut. They dissected the ganglia from the spinal
chord and we put the roots in agarose prior to osmium. They do not want
to do electron microscopy on the tissue but need the resin embedding.

The second thing they want to do is quantitate the Tol blue staining
from the thick sections. Is that doable and reproducible?

thanks,
Karen
UT Southwestern Medical Center
Live Cell and Electron Microcopy Core Facilities
214-648-2805

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:18:24 -0600
Subject: [Microscopy] viaWWW: SEM of Wood?

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Organization: yarmouk university

Title-Subject: [Filtered] sem

Message: can i insert wood in electron scanning microscope quanta 200

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From: microscopylistserver-noreply-at-microscopy.com
Date: Tue, 12 Feb 2013 22:42:54 -0600
Subject: [Microscopy] viaWWW: EMAG 2013 - Call for abstracts - deadline 22nd March 2013

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Name: Ian MacLaren

Organization: University of Glasgow / EMAG (Institute of Physics)

Title-Subject: [Filtered] EMAG 2013 - Call for abstracts - deadline 22nd
March 2013

Message: Electron Microscopy and Analysis Group Conference 2013

Organised by the IOP Electron Microscopy and Analysis Group

www.emag-iop.org

3 – 6 September 2013

University of York, York, UK

Conference highlights

High profile plenary speakers
• Prof. Ahmed Zewail (Nobel Prize in Chemistry 1999, CalTech)
• Prof. Archie Howie (University of Cambridge)
• Prof. Pratibha Gai (University of York)

Special symposium on in-situ microscopy in honour of Prof. Archie Howie

Refereed proceedings

Short courses on advanced topics in electron microscopy (Monday 2 and
Tuesday 3 September)
• Aberration-corrected electron microscopy
• Spectroscopy using electrons and X-rays in the electron microscope
• Simulation of TEM and STEM electron microscope images

Trade exhibition


Call for abstracts

We invite abstract submissions for oral or poster presentations on any
of the following topics:

Technique developments and their applications:
• The Archie Howie symposium on in-situ microscopy
• Advances in electron microscope imaging
• Spectroscopy and analysis
• Developments in STEM
• Modelling and quantification of electron microscope data
Applications of electron microscopy to specific areas:
• Nanomaterials
• Structural materials
• Functional materials
• Biomaterials and hard-soft matter interfaces in biomedical and
environmental science

Key dates

Abstract submission deadline: 22 March 2013
Early registration deadline: 2 August 2013

Abstracts should be submitted online at www.emag-iop.org

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From: wesaia-at-iastate.edu
Date: Wed, 13 Feb 2013 01:24:57 -0600
Subject: [Microscopy] viaWWW: SEM of Wood?

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There should be no problem examining wood in the Quanta as far as the SEM is concerned. The question will be what changed take place in the sample and if they are acceptable.

You might need to use environmental mode if it is essential to keep the moisture content the same. Otherwise, you could use variable pressure mode to eliminate charging without coating.

What are you looking for in your sample that you want to use SEM in the first place?

Warren Straszheim
Materials Analysis Lab, Iowa State University
home of a Quanta 250 FEG and other equipment

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Name: hadeel

Organization: yarmouk university

Title-Subject: [Filtered] sem

Message: can i insert wood in electron scanning microscope quanta 200

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22, 39 -- Subject: RE: [Microscopy] viaWWW: SEM of Wood?
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From: vitalylazar-at-att.net
Date: Wed, 13 Feb 2013 01:48:55 -0600
Subject: [Microscopy] Re: viaWWW:CCD camera sensor size

Contents Retrieved from Microscopy Listserver Archives
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Here is one:

http://www.edmundoptics.com/learning-and-support/technical/learning-center/application-notes/imaging/electronic-imaging-resource-guide/?&pagenum=5#4.3

google "CCD size" for more


Vitaly Feingold
SIA
2773 Heath Lane
Duluth GA 30096
Ph. 770-232-7785
Fax 770-232-1791
www.sia-cam.com
vitaly-at-sia-cam.com

On 2/12/2013 11:15 PM, microscopylistserver-noreply-at-microscopy.com wrote:
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} Name: Amit
}
} Organization: IISc
}
} Title-Subject: [Filtered] CCD camera sensor size
}
} Message: Good evening,
} I was wondering if anyone knows any way to find out exact sensor size of
} ccd camera? (i want to measure actual distance, like in films, between
} diffraction spots, any other suggestions are also welcome!)
} i read the brochure of manufacturer, many things dont make sense there,
} e.g chip size- 2/3 inches, 10.2mmx8.3mm
} but wont 10.3mm x 8.3 mm be 1.31 cm diagonally? while 2/3 inches should
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From: microscopylistserver-noreply-at-microscopy.com
Date: Wed, 13 Feb 2013 04:00:14 -0600
Subject: [Microscopy] viaWWW:sample holder for Baltec RES 010

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Email: ludo.rossou-at-ua.ac.be
Name: Rossou Ludo

Organization: University of Antwerp EMAT group (Belgium)

Title-Subject: [Filtered] sample holder for Baltec RES 010

Message: Dear all,

I am looking for sample holders for the old Baltec RES 010. In our lab,
a few are still working.
I think that it can be that somewhere this ion milling machines are not
in use anymore.
So I am very interested in all kind of holders.

Contact me directly.

Thanks in advance,
Ludo

Rossou Ludo
Universiteit Antwerpen
Departement Natuurkunde
Dienst EMAT
Groeneborgerlaan 171
B-2020 Antwerpen
Belgium
Tel. 00 32 3 265 32 55
Fax.00 32 3 265 33 18
ludo.rossou-at-ua.ac.be
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From: oshel1pe-at-cmich.edu
Date: Wed, 13 Feb 2013 07:08:00 -0600
Subject: [Microscopy] Re: viaWWW:Electron microscopy schools?

Contents Retrieved from Microscopy Listserver Archives
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Serene,

Central Michigan University.
We have a "microscopy major" - a microscopy option within the biology
major. The courses are light microscopy, scanning laser confocal
microscopy, transmission electron microscopy, and scanning electron
microscopy (with EDS if needed by the student). They all cover both
theory and practical lab work, and are full semester courses leading to
a B.Sc.
The courses are also open to non-majors, so we commonly have materials
science (undergrad and PhD students) and geology students..

Phil

} Email: serene_ng-at-dsi.a-star.edu.sg
} Name: Serene
}
} Title-Subject: [Filtered] Electron microscopy school
}
} Message: Dear Listers,
}
} I am looking for electron microscopy school(s) which offer short or
} full-term courses (TEM in particular) - both theory and practical
} lessons will be good.
}
} I know there's Lehigh and Hooke which offer short courses. Are there
} other institutions fellow listers can suggest? Are there similar schools
} hosted in Europe or
} Asia?
}
} Thank you listers!
} Serene

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

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From: ben.micklem-at-pharm.ox.ac.uk
Date: Wed, 13 Feb 2013 10:49:11 -0600
Subject: [Microscopy] Re: Nikon 1 series cameras on microscopes

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Back in January I posted a question about using a Nikon 1 series
inter-changeable lens, small sensor camera for microscopy.

These cameras have a 1" sensor (the recent thread on CCD sizes reminded me
to get back to the list with my experiences), which is ideal for this
application.

I bought a Nikon V1, which is heavily discounted at the moment, less than
30% of the new RRP: Ł250 with lens.
We also bought a c-mount to Nikon 1 series adapter, IR remote release,
fast SD card and card reader, mini-HDMI to HDMI cable, and a cheap full HD
monitor with HDMI input (total cost under Ł140).

So for under Ł400 GBP we have a complete imaging system.

Some photos of the set-up:

http://booking.mrc.ox.ac.uk/NikonV1_microscope_mount.jpg

We fitted this to a Leitz Dialux microscope (160mm tube length) with a 1x
c-mount adapter on the trinocular head's photo port. The C-mount adapter
gives perfect focus registration with the image in the oculars.

The shutter is fired by IR remote release. After turning on the camera, it
takes three button presses to engage the mode where it waits for the IR
signal. All other settings (e.g. Exposure mode, white balance and ISO) are
saved when the camera is turned off. The camera will only work in fully
manual exposure (the lenses normally communicate electronically with the
body, and without this, the camera isn't very helpful). The live view on
the monitor doesn't show the effect of changing the exposure, it is always
auto-gained. It does show white balance effects though. Having said that,
as it will do a quick review of the shot after it is taken, and changing
the shutter speed is just a matter of nudging a lever on the back of the
camera, if only takes a few shots to get the right exposure.

We are using the live view on a 20" LCD via HDMI, which has very fast
frame rate and is lag-free for focussing and searching.

The camera does show the imperfections in the peri-plan optics of the
low-end EF objectives, but cropping to the central region gets rid of the
soft edges.

E.g. This example taken with the 10x 0.25NA EF objective:
http://booking.mrc.ox.ac.uk/exampleV1image.jpg

The camera can also taken impressive full HD movies at 60fps, so would be
useful for teaching purposes.

Hope this mini-review is of use to someone looking for colour brightfield
imaging on a low budget.

Ben

--
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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From: WHITTAKS-at-si.edu
Date: Wed, 13 Feb 2013 14:49:01 -0600
Subject: [Microscopy] =?iso-8859-1?Q?=B5CT_on_the_SEM?=

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html


Anyone out there using the SEM mounted microCT scanners? Would like to discuss real world practicality and use examples as my Museum considers the future of CT going forward and this might solve a dilemma we are having in addressing the range of sample sizes and resolutions.


Scott Whittaker
Manager SEM Lab
Smithsonian Institution
National Museum of Natural History
PO Box 37012   MRC104
Washington DC 20013-7012
202-633-0891




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From: Naomi_McCallum-at-health.qld.gov.au
Date: Wed, 13 Feb 2013 20:33:12 -0600
Subject: [Microscopy] Fwd: Re: viaWWW:CCD camera sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi Amit

Forgive me if I have missed something obvious since I only have experience with biological specimens and 1 vendor's software.

My understanding is that if the images is calibrated then the distance can be calculated from the image using software. The number of pixels x the distance per pixel gives the measured distance: measurement software can do this for you, you just select the 2 points. If the images you speak of is acquired on a CCD camera, then it should be (or can be) calibrated, and the camera software may allow the measurements you require.

If you are attempting to measure distance on an image that has been exported from that software since acquisition then care should be taken since you may not have the original correct calibration information associated with the image. Keep in mind such things as binning, changes to pixel size/shape etc on post processed images.

There are others on the list with more experience but hope this helps.
Naomi

} } } {vitalylazar-at-att.net} 2/13/2013 5:53 pm } } }



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Here is one:

http://www.edmundoptics.com/learning-and-support/technical/learning-center/application-notes/imaging/electronic-imaging-resource-guide/?&pagenum=5#4.3

google "CCD size" for more


Vitaly Feingold
SIA
2773 Heath Lane
Duluth GA 30096
Ph. 770-232-7785
Fax 770-232-1791
www.sia-cam.com
vitaly-at-sia-cam.com

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} Title-Subject: [Filtered] CCD camera sensor size
}
} Message: Good evening,
} I was wondering if anyone knows any way to find out exact sensor size of
} ccd camera? (i want to measure actual distance, like in films, between
} diffraction spots, any other suggestions are also welcome!)
} i read the brochure of manufacturer, many things dont make sense there,
} e.g chip size- 2/3 inches, 10.2mmx8.3mm
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From: Naomi_McCallum-at-health.qld.gov.au
Date: Wed, 13 Feb 2013 22:14:25 -0600
Subject: [Microscopy] Re: viaWWW:microtome accuracy

Contents Retrieved from Microscopy Listserver Archives
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Hi Judy

Unfortunately we don't have much detail on what (size) you will be attempting to measure and thus degree of accuracy that you are seeking. However, even with the "most accurate" microtome on the market, a lot can happen from tissue dissection through processing/freezing, and especially during & after microtomy that can affect accuracy .

Here's an idea from left field: a researcher engaging our services injected fluorescent polystyrene beads (commercial name escapes me at present) into their specimens to locate area of interest. (Excellent for accurate sampling and dissection.) When processed, the beads dissolved but left replicas as perfect resin circles within the tissue (ie in the fat adjacent to the area of interest.

If this was applied to frozen section, the beads would not dissolve if aqueously mounted, and the fluorescent label would allow localisation of the beads for measurement calibration. Or if paraffin processing was employed it would be the same as we experienced with the resin, you have to look for the regular spherical spaces where the beads once were.

If this is not helpful, feel free to file appropriately.

Kind regards
Naomi

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Email: trogadisj-at-smh.ca
Name: Judy Trogadis

Organization: St. Michael's Hospital

Title-Subject: [Filtered] microtome accuracy

Message: Fellow microscopists

We are planning to do very fine optical measurements from tissue
sections, therefore, control of thickness and of even surface is critical.

Which are the most accurate and reproducible microtomes on the market
for paraffin or frozen blocks? Are there laser based tissue sectioning
equipment.

Any suggestions are welcomed, including replies from manufacturers.
Thank you,
Judy


Judy Trogadis
Bio-Imaging Coordinator
Keenan Research Centre, St. Michael's
209 Victoria Street
Toronto M5B 1T8, Canada
office: 416-864-6060 ext. 77612
imaging facility: ext. 77434
cell: 416-254-9330
trogadisj-at-smh.ca


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From: microscopylistserver-noreply-at-microscopy.com
Date: Thu, 14 Feb 2013 01:20:51 -0600
Subject: [Microscopy] viaWWW:microscopy outreach - microscopes

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Email: kalennon-at-hagerstowncc.edu
Name: Kristen Lennon

Organization: Hagerstown Community College

Title-Subject: [Filtered] microscopy outreach - microscopes

Message: Hello Everyone,

I've just been given the exciting and daunting challenge of creating a
K-12 outreach and teacher education program centered on microscopy.I've
visited the Project Micro web site and am using those resources, but I'm
wondering if any of you have any other suggested resources. I'd also
like some suggestions as to where to buy good quality stereo microscopes
at a K-12 non-profit kind of price. The idea is to get more kids excited
about science through the engaging microscopic world and make microscopy
more accessible to K-12 students and their teachers.

Your advice is, as always, much appreciated.
Best,
Kristen Lennon
kalennon-at-hagerstowncc.edu

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From: amit.welcomes.u-at-gmail.com
Date: Thu, 14 Feb 2013 03:27:36 -0600
Subject: [Microscopy] CCD camera sensor size

Contents Retrieved from Microscopy Listserver Archives
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Thank you for replying first of all.
Thank you Vitaly Feingold for the link it helped a lot in clearing up
jargon. Though no solutions were given!

Naomi McCallum: Actually i want to measure varios constants of microscope
for example exact magnification and spherical aberration etc. for that what
we require is how much the image feature has actually "moved". earlier when
films were used it was easy as you can take out the films and measure it
with a foot ruler or something(thats what is sounds like from papers i have
read, if i am missing something then sorry, may be someone more experienced
might correct me). but to find out that now i need what is size of each
pixel.
software allots size of each pixel according to values given to it (eg.0466
nm /pixel etc). but actual pixel size is in micrometers, i.e physical size
of sensor on camera. To avoid binning etc i am making sure that camera runs
on maximum resolution.


I have tried following method but dont know if its correct.-
1. obtain diffraction pattern of some single crystal sample
2.increase the camera length to magnify it
3.fluorescent screens have markers on them (jeol one that we are using has
5mm ticks at the lower side)
4. move any spot on that tticks and take images.
5. you can get physical distance from that ticks(ie. 5mm) and number of
pixels moved in image.
we can then find out whats the exact physical size of pixel

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From: benoit.zuber.work-at-gmail.com
Date: Thu, 14 Feb 2013 04:24:01 -0600
Subject: [Microscopy] CEMOVIS/frozen-hydrated sections course announcement

Contents Retrieved from Microscopy Listserver Archives
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Dear all,

We are pleased to announce

the 3rd UB Practical Course on Cryo-Electron Microscopy of Vitreous
Sections (CEMOVIS, a.k.a cryoEM of frozen-hydrated sections)

2-4 July 2013 / 9-11 July 2013

Organiser : Benoît Zuber
Instructors: Benoît Zuber, Daniel Studer, Ioan Iacovache

The objective of the course is to teach participants the practical
skills necessary to successfully apply CEMOVIS (a.k.a. Frozen-hydrated
sections) in their laboratories. Our latest tricks making CEMOVIS
quite easy will be demonstrated. Essential background theory of
CEMOVIS will be given and most of the time will be spent practicing
high-pressure freezing, cryo-sectioning, and low-dose TEM imaging of
vitreous sections. 1 high-pressure freezing machine, 3
state-of-the-art cryo-ultramicrotomes and 1 cryo-electron microscope
will be dedicated to the course. The 3-day course is given twice for
two different groups of maximum 4 participants each so that every
participant can be actively practicing during the whole course.

All the participants of the previous previous courses have been
successful in producing, collecting and imaging vitreous sections.

The course is intended for scientists whose research projects will
benefit from the use of CEMOVIS. Experience in either cryo-electron
microscopy or ultramicrotomy of resin-embedded specimens is a
prerequisite.

Information and registration:
http://www.ana.unibe.ch/events/cemovis/index.html

Registration deadline: 17.03.2013

Yours faithfully
Benoît Zuber
Note: do not reply to this e-mail address but to the one in my signature below.
--
Prof. Benoît Zuber
Institute of Anatomy
University of Bern
Baltzerstrasse 2
3000 Bern 9
Switzerland
benoit.zuber-at-ana.unibe.ch
+41 31 631 84 40


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From: ben.micklem-at-pharm.ox.ac.uk
Date: Thu, 14 Feb 2013 05:33:30 -0600
Subject: [Microscopy] Re: CCD camera sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I think you may be confusing with the physical dimensions of the photosite
on the CCD, and the dimensions of the image projected onto one photosite-
which leads to one pixel in the image.

The camera manufacturer should state the size of the photosites in the
spec sheet, if they don't, you can divide the horizontal or vertical
distance by the number of photosites/pixels in that axis.


It that it sounds like you are using a TEM, so in this case there is an
added complexity: (unless you are using a fancy direct electron capture
CCD) the electrons hit a scintillator, this leads to photons being
emitted, and these are then then either reflected off a mirror into a
lens, or down optic fibres to the CCD. The size of the image created by
the electrons on the scitillator may not be the same size as the photon
image reaching the CCD. Also, the electron image is getting larger as you
lower plane of the scintillator. The camera length will be calculated to
the location of the normal view screen, so if you are using that for you
scaling calulations, you have to take into account how the camera length
will be changed by the location of the scintillator plane (if the
scintintillator is say 10cm below the screen, this will not mean the
camera length is 10cm greater! It will require more thought, and possibly
knowing how hight above the view screen and the camera's scintillator the
nodal point of the projector lens is? But my electron optical knowledge is
lacking, sadly).

I have never worked with diffraction images (I am a biologist), and I
don't know what the exact relationship between camera length and
magnification is, or whether diffraction images really have a
'magnification' as such- I am completely ignorant!

Hope that helps you (even if it makes things more confusing in the short
term!),

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 14/02/2013 09:33, "amit.welcomes.u-at-gmail.com"
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From: schooley-at-mcn.org
Date: Thu, 14 Feb 2013 16:14:03 -0600
Subject: [Microscopy] Microscopy outreach - microscopes

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Kristen -

Project MICRO will be delighted to help you as much as we can.
You're already working with MICRO's website, so I'll avoid
duplicating its contents; I'll restrict myself to your specific
concerns.

Teacher's manuals. "Microscopic Explorations" was produced as part of
the Lawrence Hall of Science's Great Explorations in Math and Science
(GEMS) series, with sponsorship by MSA; it's middle school. "Private
Eye" is an independently produced K-12 manual with a different
approach. "Nanoscale Science" is a NSTA manual for grades 6-12.
You'll find full information on the first two in MICRO' main
booklist; "Nanoscale Science" is in the 'Nanotechnology for Kids"
list. A dozen articles about various aspects of microscopy outreach
have appeared recently in MSA's 'Microscopy Today' magazine. You
can read them online; they're listed in the introduction to the MICRO
booklist.

Online lesson. Lesley Bechtold {Lesley.Bechtold-at-jax.org} of the
Jackson Lab in Bar Harbor, Maine has written an online microscopy
lesson that MICRO hopes to offer soon; you might be able to use it.
Contact Lesley directly for information.

Teacher education. Both GEMS and Private Eye offer excellent onsite
teacher training programs.

Microscopes. You say "stereo". That just means "two eyepieces". Do
you mean low power inspection/dissection scopes, compound scopes, or
both? I've got a lot to say about buying school microscopes on the
website, and none of it encourages binocular optics. Monocular is
much cheaper. Rough use is inevitable, and repairing misaligned
optics requires professional help. An estimated 17% of younger
children have some sort of binocular vision problem, and young faces
haven't the eye spacing to use many binocs.

Having said that, my basic advice is that you should work with a
company that specializes in student microscopes, rather than a
general school supply or lab equipment firm. All of the scopes are
Chinese, and they're all quite similar; the difference will be in
quality control and repair service. If you'd like help with onsite
evaluation of samples, MICRO can find an experienced MSA member in
your area.

I welcome your further questions!

Caroline

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Email: kalennon-at-hagerstowncc.edu
Name: Kristen Lennon

Organization: Hagerstown Community College

Title-Subject: [Filtered] microscopy outreach - microscopes

Message: Hello Everyone,

I've just been given the exciting and daunting challenge of creating a
K-12 outreach and teacher education program centered on microscopy.I've
visited the Project Micro web site and am using those resources, but I'm
wondering if any of you have any other suggested resources. I'd also
like some suggestions as to where to buy good quality stereo microscopes
at a K-12 non-profit kind of price. The idea is to get more kids excited
about science through the engaging microscopic world and make microscopy
more accessible to K-12 students and their teachers.

Your advice is, as always, much appreciated.
Best,
Kristen Lennon
kalennon-at-hagerstowncc.edu

--
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

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From: wa5ekh-at-juno.com
Date: Fri, 15 Feb 2013 21:34:07 -0600
Subject: [Microscopy] Basic Microscope Questions-

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

There are a few things I am curious about:

Concerning a recent statement.."A monocular microscope is much cheaper.", I started thinking (dangerous!).

Being a user, not a microscope optics technologist, I always had several questions:

1)Don't Binocular microscopes simply split the image? What exactly is the advantage(s)in a compound turret-type microscope)? Isn't this an obsolete function and design?(Pre-electronic camera)?

2)Also, it would seem 'logical' to use ccd cameras and monocular microscopes.
a)since there are fewer optical components in monocular, so less optical distortions(minimal absorption, refraction, diffraction,..?, right?
b) and 'how much' information is lost or degraded by electronic amplification, scanning or dithering(?) artifacts and CCD noise(& etc.)? Are the losses reduced by higher ccd pixel resolution? I have some confusion about CCDs, and I would guess that I am not alone. I suppose the higher number of pixels advertised or stated in CCD specs. is actually suggesting a higher density per square inch..is this equivalent or similar to 'spatial resolution'(and how is it comparable)?

3)And finally, which produces more usable information: monocular, binocular or trinocular microscopes(these more advanced levels would seem to require many more optical components).

(I think this is just the tip of the iceberg..is there a very thorough website for these details..seems like I remember someone mentioning a few...frequently)
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From: amit.welcomes.u-at-gmail.com
Date: Mon, 18 Feb 2013 05:04:53 -0600
Subject: [Microscopy] CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

The "recent statement" was mine, in response to a question about
school microscopes. I'll respond to your questions from that point
of view.
1) Some dissecting scopes have 2 fully separate light paths,
providing a true stereo image.; the cheaper ones do not. Such scopes
are used to look at the surface of a sample, so stereo is useful.
Compound scopes are used with transmitted light to view the internal
structure of thin samples. Obsolete? Perhaps eventually, but at the
K-12 level, cost is still in control.
2a) The 2 light paths are parallel rather than in series, so the
defects aren't additive.
2b) Not relevant for basic school microscopes.
3) Silly question. The 3rd tube is just a place to mount a camera
or other device.
The website that you remember is Molecular Expressions; it's very
good. http://micro.magnet.fsu.edu/primer/

Caroline

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--
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

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Sorry for late reply everyone. thanks for the responses!
-at-Ben- yes you are right i want the physical size of the photosite
which makes one pixel on image. as from it i can actually count
movement in cm etc by measuring pixcels.

Sadly its not a Gatan camera. And procuring a standard sample like
mag*i*cal will not be feasible, vis a vis money and time.


Regards
Amit Gupta

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From: wesaia-at-iastate.edu
Date: Mon, 18 Feb 2013 08:37:53 -0600
Subject: [Microscopy] CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Maybe I am missing something here. Correct me if I am. Back in the old days, I knew the size of my Polaroid film exactly. That would be like knowing the size of your sensor (or its pixels) exactly. Your question is about the actual magnification.

Do not trust the magnification display on your scope. I have heard that +/- 5% errors are not uncommon. Most systems are better. You need to calibrate the scope under the same conditions that you are using for your sample.

I understand that one of the benefits of Mag*i*cal is that it provides several functions and scales on the same sample so that it is useful at multiple magnifications.

However, the standard may not have to be exotic. You might use a TEM grid and count the grid openings. You can then determine your pixel spacing from the number of pixels across a feature of know dimension. You should also be able to determine the magnification. Just don't assume that the displayed magnification is correct.

Warren S.

-----Original Message-----
X-from: amit.welcomes.u-at-gmail.com [mailto:amit.welcomes.u-at-gmail.com]
Sent: Monday, February 18, 2013 5:06 AM
To: wesaia-at-iastate.edu

Sorry for late reply everyone. thanks for the responses!
-at-Ben- yes you are right i want the physical size of the photosite
which makes one pixel on image. as from it i can actually count
movement in cm etc by measuring pixcels.

Sadly its not a Gatan camera. And procuring a standard sample like
mag*i*cal will not be feasible, vis a vis money and time.


Regards
Amit Gupta

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From: dmitry.v.sokolov-at-gmail.com
Date: Mon, 18 Feb 2013 15:31:43 -0600
Subject: [Microscopy] CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
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The sensor is a part of the optical / detection system. As an image is the output of the instrument, the sensor size creates confusion in the image calibration and can be excluded from calculations:
http://confocal-manawatu.pbworks.com/w/page/60817289/Image%20Calibration

Just try to avoid calibration of your equipment and images by pixel size or scale bar.

Cheers,
Dmitry
Advanced Knowledge Management
for MICROSCOPY and Image Analysis
_________________________________________
Dmitry Sokolov, Ph.D.
Mobile: +64 21 063 5382
dmitry.v.sokolov-at-gmail.com

On 19/02/2013 3:48 a.m., wesaia-at-iastate.edu wrote:
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} Maybe I am missing something here. Correct me if I am. Back in the old days, I knew the size of my Polaroid film exactly. That would be like knowing the size of your sensor (or its pixels) exactly. Your question is about the actual magnification.
}
} Do not trust the magnification display on your scope. I have heard that +/- 5% errors are not uncommon. Most systems are better. You need to calibrate the scope under the same conditions that you are using for your sample.
}
} I understand that one of the benefits of Mag*i*cal is that it provides several functions and scales on the same sample so that it is useful at multiple magnifications.
}
} However, the standard may not have to be exotic. You might use a TEM grid and count the grid openings. You can then determine your pixel spacing from the number of pixels across a feature of know dimension. You should also be able to determine the magnification. Just don't assume that the displayed magnification is correct.
}
} Warren S.
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} To: wesaia-at-iastate.edu
} Subject: [Microscopy] CCD sensor size
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} Sorry for late reply everyone. thanks for the responses!
} -at-Ben- yes you are right i want the physical size of the photosite
} which makes one pixel on image. as from it i can actually count
} movement in cm etc by measuring pixcels.
}
} Sadly its not a Gatan camera. And procuring a standard sample like
} mag*i*cal will not be feasible, vis a vis money and time.
}
}
} Regards
} Amit Gupta
}
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From: mike.bode-at-resaltatech.com
Date: Mon, 18 Feb 2013 21:43:08 -0600
Subject: [Microscopy] RE: CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
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Hello Amit,

I have been following your posts, and I am not sure I understand why you
need the actual pixel size. If you want to measure diffraction spots or
their position, all you need to do is to insert a specimen with a known
crystal lattice (for example a piece of Si), and calibrate your system for
your camera length. The rest is then just a bit of arithmetic. And the
centers of the diffraction spots should be as accurate as you can get.

The same is true if you want to measure distances in real space. Insert a
calibration sample (for example the ubiquitous line grating), and measure
the distance between lines and use a simple program that allows you to get
the pixel coordinates, and you can calculate a calibration in microns/pixel.
And then you can calculate distances or movements.

Any microscopy software should be able to do this.

Am I missing something?

Can you tell us who the manufacturer of your camera is, or what the model
is? Perhaps there is a way to find the sensor that was used in the camera.

Mike

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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Sorry for late reply everyone. thanks for the responses!
-at-Ben- yes you are right i want the physical size of the photosite which
makes one pixel on image. as from it i can actually count movement in cm etc
by measuring pixcels.

Sadly its not a Gatan camera. And procuring a standard sample like mag*i*cal
will not be feasible, vis a vis money and time.


Regards
Amit Gupta

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From: amit.welcomes.u-at-gmail.com
Date: Mon, 18 Feb 2013 22:55:57 -0600
Subject: [Microscopy] CCD sensor size

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-at- Warren S.
exactly! I need to calibrate my whole TEM. From magnification to
camera length to aberration constants. for that i need what is the
size of my "polaroid film". since we have a ccd camera instead of
film, I need to know the actual size of ccd sensor (will I not?)

-at- rest (sorry for not addressing individually)
I can caliberate at high mag with gold lattice image, at low mag with
copper grid etc, its the range 80000 to 200000 which is giving bit
trouble.

camera: Olympus Keen View G2

Gatan generally gives a caliberation sample with CCD, alas! olympus dont.

With Regards
Amit

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From: dmitry.v.sokolov-at-gmail.com
Date: Tue, 19 Feb 2013 00:27:26 -0600
Subject: [Microscopy] Re: CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hi Amit,

the trick with the microscopes is that they can be considered as a
single "filter" between your sample and your computer screen / eye.

Calibrate the instrument against known (better if with the standard
reference) sample at the magnification and other critical parameters
fixed. The subsequent calibration of the image is a breeze:
Image Size (um) = Maximum Visible Length of Reference (um) * Image
Size (pixels) / Maximum Visible Length of Reference (pixels)

The pixel size at given (same as at the calibration of the instrument!)
imaging conditions is calculated as:
Image Size (um) / Image Size (pixels).

Estimation of the camera pixel size:
Camera pixel size (um) = Maximum Visible Length of Reference (um) /
Maximum Visible Length of Reference (pixels) / Magnification

Here we come to the need of calibration of the microscope in terms of
magnification at the plane of the camera chip that may be tricky. I
would rely in this case on either chip data from the manufacturer of the
camera or direct measurements of the chip size under the binocular
microscope. Don't forget about the LM calibration too. ;0)

This all is described at MIAWiki Knowledge Network:
http://confocal-manawatu.pbworks.com/w/page/60817289/Image%20Calibration
If something is missing or unclear, your comments/suggestions at the
bottom of the page will be highly appreciated.

Please Skype me: FalconDot if I can be of more help in real time.

Cheers,
Dmitry

Advanced Knowledge Management
for MICROSCOPY and Image Analysis
________________________________
Dmitry Sokolov, Ph.D.
Mob: +64 21 063 5382
dmitry.v.sokolov-at-gmail.com

19.02.2013 18:05, amit.welcomes.u-at-gmail.com ďčřĺň:
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} ----------------------------------------------------------------------------
}
} -at- Warren S.
} exactly! I need to calibrate my whole TEM. From magnification to
} camera length to aberration constants. for that i need what is the
} size of my "polaroid film". since we have a ccd camera instead of
} film, I need to know the actual size of ccd sensor (will I not?)
}
} -at- rest (sorry for not addressing individually)
} I can caliberate at high mag with gold lattice image, at low mag with
} copper grid etc, its the range 80000 to 200000 which is giving bit
} trouble.
}
} camera: Olympus Keen View G2
}
} Gatan generally gives a caliberation sample with CCD, alas! olympus dont.
}
} With Regards
} Amit
}
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From: larry.stoter-at-gmail.com
Date: Tue, 19 Feb 2013 01:00:39 -0600
Subject: [Microscopy] Re: CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Amit,

For magnifications where you cannot directly use a standard to calibrate, you can use ratios.

So, for example, starting from the lowest high-magnification where you can calibrate directly from a lattice image, find a region on the sample with two clearly visible and widely separated features - measure the spacing. Drop the magnification and re-measure the spacing. The ratio of the two measurements gives you the ratio of the lower, uncalibrated magnification to the higher, calibrated magnification.

Larry Stoter
(Working on a Microsoft-free computer)

On 19 Feb 2013, at 05:04, amit.welcomes.u-at-gmail.com wrote:

}
}
}
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} ----------------------------------------------------------------------------
}
} -at- Warren S.
} exactly! I need to calibrate my whole TEM. From magnification to
} camera length to aberration constants. for that i need what is the
} size of my "polaroid film". since we have a ccd camera instead of
} film, I need to know the actual size of ccd sensor (will I not?)
}
} -at- rest (sorry for not addressing individually)
} I can caliberate at high mag with gold lattice image, at low mag with
} copper grid etc, its the range 80000 to 200000 which is giving bit
} trouble.
}
} camera: Olympus Keen View G2
}
} Gatan generally gives a caliberation sample with CCD, alas! olympus dont.
}
} With Regards
} Amit
}
} ==============================Original Headers==============================
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From: mike.bode-at-resaltatech.com
Date: Tue, 19 Feb 2013 09:17:20 -0600
Subject: [Microscopy] CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Hello Amit,

Can you give me the serial number of the camera? I can then get you the
pixel size of the camera.

For a magnification of 8000 - 20,000 the Diffraction Grating Replica,
available from a number of vendors, should give you what you need to
calibrate your system.

If you look on the DVD or CD for the software that came with your camera,
you should find a "step-by-step" document (in the doc folder), which
explains the calibration routine for the cameras. It is pretty straight
forward. You can store a number of calibration values for different
magnifications, and the software (iTEM) will interpolate between the
calibrated values. Let me know if you found the document. I can also send it
to you, but I do need to know what software and what release you are using.

Can you tell me where the TEM is located?

Mike



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 Message-----
X-from: amit.welcomes.u-at-gmail.com [mailto:amit.welcomes.u-at-gmail.com]
Sent: Monday, February 18, 2013 10:03 PM
To: mike.bode-at-resaltatech.com

-at- Warren S.
exactly! I need to calibrate my whole TEM. From magnification to camera
length to aberration constants. for that i need what is the size of my
"polaroid film". since we have a ccd camera instead of film, I need to know
the actual size of ccd sensor (will I not?)

-at- rest (sorry for not addressing individually) I can caliberate at high mag
with gold lattice image, at low mag with copper grid etc, its the range
80000 to 200000 which is giving bit trouble.

camera: Olympus Keen View G2

Gatan generally gives a caliberation sample with CCD, alas! olympus dont.

With Regards
Amit

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From: Edelmare-at-miamioh.edu
Date: Thu, 21 Feb 2013 10:43:43 -0600
Subject: [Microscopy] Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear All,

Does anyone have any step-by-step notes they're willing to share on how to
use the strain mapping software within ESVision (including screen shots?)?

I have some handwritten notes taken a few years ago, but they seem to make
little sense with the version of ESVision I am using (Version 5.0).
Likewise, the Help menu on ESVision is hopeless- there is no mention of it
anywhere and certainly nothing on how to set it up.

I'd be interested to hear other recommendations on getting strain maps out
of Emispec files.

Yours, Jon

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Hello listers, got an odd question that I am hoping the more experienced
microanalytical folks can help me with.

I think we are seeing some type of "Escape" peak using an SDD XEDS
system, but these are not the 1.740kev Silicon escape peaks I am used to.
They are much closer to the primary peak, vary in energy displacement from
the primary peaks, and seem to be proportionally larger than Silicon escape
peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at- 4.508kev),
0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).

They are not "system peaks" as they track with the larger primary peaks as
we change samples. I suspect that they are "escape artifacts" of SDD´s and
obviously I do not spend enough time reading "MicroNews" and the
microanalysis literature to be aware of them. Anyone want to help us out,
please?

We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps (TEM
remember?), but have tested with same results at higher 10 - 60kcps.

The Bruker software does not provide any identification markers. As they
are a significant size, if they are an escape artifact of SDD´s, their absence
from the primary peak will significantly effect the quant calculations. Is this
correct?


Thank you in advance!


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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From: wesaia-at-iastate.edu
Date: Thu, 21 Feb 2013 11:19:02 -0600
Subject: [Microscopy] Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I wonder if it might be some sort of incomplete charge collection or else a calibration error. We ran into incomplete charge collection on a broken Ge detector. It broke after it warmed up and we got a shadowing of the spectrum downscale from where it was suppose to be. As I recall, we found peaks at about 60% of the energy of the main peaks. Yours seems to be consistently around 92% of the energy of the main peak (i.e., ~8% downscale).

I mention calibration error in case different portions of the detector feed into different preamps. (Does anyone do that?) Maybe one segment is out of calibration.

I hope your system is under warranty. I would guess you need to get Bruker in to evaluate the detector and/or its setup. Maybe it is just a calibration issue.

I would be interested in seeing some of the data if you could provide a screen shot or an MSA copy of a spectrum.

Warren

-----Original Message-----
X-from: Edelmare-at-miamioh.edu [mailto:Edelmare-at-miamioh.edu]
Sent: Thursday, February 21, 2013 10:45 AM
To: wesaia-at-iastate.edu


Hello listers, got an odd question that I am hoping the more experienced
microanalytical folks can help me with.

I think we are seeing some type of "Escape" peak using an SDD XEDS
system, but these are not the 1.740kev Silicon escape peaks I am used to.
They are much closer to the primary peak, vary in energy displacement from
the primary peaks, and seem to be proportionally larger than Silicon escape
peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at- 4.508kev),
0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).

They are not "system peaks" as they track with the larger primary peaks as
we change samples. I suspect that they are "escape artifacts" of SDD´s and
obviously I do not spend enough time reading "MicroNews" and the
microanalysis literature to be aware of them. Anyone want to help us out,
please?

We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps (TEM
remember?), but have tested with same results at higher 10 - 60kcps.

The Bruker software does not provide any identification markers. As they
are a significant size, if they are an escape artifact of SDD´s, their absence
from the primary peak will significantly effect the quant calculations. Is this
correct?


Thank you in advance!


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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From: Edelmare-at-miamioh.edu
Date: Thu, 21 Feb 2013 12:00:24 -0600
Subject: [Microscopy] Unidentified XEDS Peak - Example image

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html



O.k., well thank you folks for thinking about it.

Here is an example of what we are seeing:

http://www.cami.muohio.edu/xeds/Weirdpeak1.jpg


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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From: Edelmare-at-miamioh.edu
Date: Thu, 21 Feb 2013 12:02:25 -0600
Subject: [Microscopy] RE: Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Warren:

Not a calibration error, as the peaks above and below all align where they
should. Hmm, but I hope its not a "shadowing as you suggest.

Image at:

http://www.cami.muohio.edu/xeds/Weirdpeak1.jpg




On 21 Feb 2013 at 17:18, Straszheim, Warren E [BIOTC] wrote:

} I wonder if it might be some sort of incomplete charge collection or
} else a calibration error. We ran into incomplete charge collection on
} a broken Ge detector. It broke after it warmed up and we got a
} shadowing of the spectrum downscale from where it was suppose to be.
} As I recall, we found peaks at about 60% of the energy of the main
} peaks. Yours seems to be consistently around 92% of the energy of the
} main peak (i.e., ~8% downscale).
}
} I mention calibration error in case different portions of the detector
} feed into different preamps. (Does anyone do that?) Maybe one segment
} is out of calibration.
}
} I hope your system is under warranty. I would guess you need to get
} Bruker in to evaluate the detector and/or its setup. Maybe it is just
} a calibration issue.
}
} I would be interested in seeing some of the data if you could provide
} a screen shot or an MSA copy of a spectrum.
}
} Warren
}
} -----Original Message-----
} From: Edelmare-at-miamioh.edu [mailto:Edelmare-at-miamioh.edu]
} Sent: Thursday, February 21, 2013 10:45 AM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] Subject: Unidentified extra XEDS peaks
}
}
}
}
} ----------------------------------------------------------------------
} ------ The Microscopy ListServer -- CoSponsor: The Microscopy Society
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} http://www.microscopy.com/MicroscopyListserver On-Line Help
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} ----------------------------------------------------------------------
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}
} Hello listers, got an odd question that I am hoping the more
} experienced microanalytical folks can help me with.
}
} I think we are seeing some type of "Escape" peak using an SDD XEDS
} system, but these are not the 1.740kev Silicon escape peaks I am used
} to. They are much closer to the primary peak, vary in energy
} displacement from the primary peaks, and seem to be proportionally
} larger than Silicon escape peaks from Si\Li detectors. Examples:
} 0.372 kev below Ti Ka (-at- 4.508kev), 0.610kev below Ni Ka (-at- 7.471
} kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
}
} They are not "system peaks" as they track with the larger primary
} peaks as we change samples. I suspect that they are "escape
} artifacts" of SDD´s and obviously I do not spend enough time reading
} "MicroNews" and the microanalysis literature to be aware of them.
} Anyone want to help us out, please?
}
} We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps
} (TEM remember?), but have tested with same results at higher 10 -
} 60kcps.
}
} The Bruker software does not provide any identification markers. As
} they are a significant size, if they are an escape artifact of SDD´s,
} their absence from the primary peak will significantly effect the
} quant calculations. Is this correct?
}
}
} Thank you in advance!
}
}
} 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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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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From: milesd-at-us.ibm.com
Date: Thu, 21 Feb 2013 14:40:29 -0600
Subject: [Microscopy] Re: Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

I am by no means very knowledgeable in the subject, but isn't that a sum
peak? ( Ti Ka1+2)

Regards,
Darrell


Edelmare-at-miamioh.edu wrote on 02/21/2013 11:44:39 AM:

} From: Edelmare-at-miamioh.edu
} To: Darrell Miles/Fishkill/IBM-at-IBMUS,
} Date: 02/21/2013 11:55 AM
} Subject: [Microscopy] Subject: Unidentified extra XEDS peaks
}
}
}
}
}
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America
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}
} Hello listers, got an odd question that I am hoping the more experienced

} microanalytical folks can help me with.
}
} I think we are seeing some type of "Escape" peak using an SDD XEDS
} system, but these are not the 1.740kev Silicon escape peaks I am used
to.
} They are much closer to the primary peak, vary in energy displacement
from
} the primary peaks, and seem to be proportionally larger than Silicon
escape
} peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at-
4.508kev),
} 0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
}
} They are not "system peaks" as they track with the larger primary peaks
as
} we change samples. I suspect that they are "escape artifacts" of SDD´s
and
} obviously I do not spend enough time reading "MicroNews" and the
} microanalysis literature to be aware of them. Anyone want to help us
out,
} please?
}
} We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps (TEM
} remember?), but have tested with same results at higher 10 - 60kcps.
}
} The Bruker software does not provide any identification markers. As
they
} are a significant size, if they are an escape artifact of SDD´s,
} their absence
} from the primary peak will significantly effect the quant
} calculations. Is this
} correct?
}
}
} Thank you in advance!
}
}
} 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
}
}
}
} ==============================Original
Headers==============================
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From: klivi-at-jhu.edu
Date: Thu, 21 Feb 2013 14:53:52 -0600
Subject: [Microscopy] Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Actually, Warren's idea is compelling, even though I have not heard of this before. The energy ratio of the "shadow" peak to the characteristic peak is constant at 0.918+-0.01. Sounds like there are two calibrations acting here with different gains.
Ken

On Feb 21, 2013, at 1:04 PM, Edelmare-at-miamioh.edu wrote:

}
}
}
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} Warren:
}
} Not a calibration error, as the peaks above and below all align where they
} should. Hmm, but I hope its not a "shadowing as you suggest.
}
} Image at:
}
} http://www.cami.muohio.edu/xeds/Weirdpeak1.jpg
}
}
}
}
} On 21 Feb 2013 at 17:18, Straszheim, Warren E [BIOTC] wrote:
}
} } I wonder if it might be some sort of incomplete charge collection or
} } else a calibration error. We ran into incomplete charge collection on
} } a broken Ge detector. It broke after it warmed up and we got a
} } shadowing of the spectrum downscale from where it was suppose to be.
} } As I recall, we found peaks at about 60% of the energy of the main
} } peaks. Yours seems to be consistently around 92% of the energy of the
} } main peak (i.e., ~8% downscale).
} }
} } I mention calibration error in case different portions of the detector
} } feed into different preamps. (Does anyone do that?) Maybe one segment
} } is out of calibration.
} }
} } I hope your system is under warranty. I would guess you need to get
} } Bruker in to evaluate the detector and/or its setup. Maybe it is just
} } a calibration issue.
} }
} } I would be interested in seeing some of the data if you could provide
} } a screen shot or an MSA copy of a spectrum.
} }
} } Warren
} }
} } -----Original Message-----
} } From: Edelmare-at-miamioh.edu [mailto:Edelmare-at-miamioh.edu]
} } Sent: Thursday, February 21, 2013 10:45 AM
} } To: wesaia-at-iastate.edu
} } Subject: [Microscopy] Subject: Unidentified extra XEDS peaks
} }
} }
} }
} }
} } ----------------------------------------------------------------------
} } ------ The Microscopy ListServer -- CoSponsor: The Microscopy Society
} } of America To Subscribe/Unsubscribe --
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} } http://www.microscopy.com/MicroscopyListserver/FAQ.html
} } ----------------------------------------------------------------------
} } ------
} }
} }
} } Hello listers, got an odd question that I am hoping the more
} } experienced microanalytical folks can help me with.
} }
} } I think we are seeing some type of "Escape" peak using an SDD XEDS
} } system, but these are not the 1.740kev Silicon escape peaks I am used
} } to. They are much closer to the primary peak, vary in energy
} } displacement from the primary peaks, and seem to be proportionally
} } larger than Silicon escape peaks from Si\Li detectors. Examples:
} } 0.372 kev below Ti Ka (-at- 4.508kev), 0.610kev below Ni Ka (-at- 7.471
} } kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
} }
} } They are not "system peaks" as they track with the larger primary
} } peaks as we change samples. I suspect that they are "escape
} } artifacts" of SDD´s and obviously I do not spend enough time reading
} } "MicroNews" and the microanalysis literature to be aware of them.
} } Anyone want to help us out, please?
} }
} } We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} } 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps
} } (TEM remember?), but have tested with same results at higher 10 -
} } 60kcps.
} }
} } The Bruker software does not provide any identification markers. As
} } they are a significant size, if they are an escape artifact of SDD´s,
} } their absence from the primary peak will significantly effect the
} } quant calculations. Is this correct?
} }
} }
} } Thank you in advance!
} }
} }
} } 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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} Center for Advanced Microscopy & Imaging
} 9C Upham Hall
} Miami University, Oxford, OH 45056
} Ph: 513.529.5712 Fax: 513.529.4243
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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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From: milesd-at-us.ibm.com
Date: Thu, 21 Feb 2013 16:34:06 -0600
Subject: [Microscopy] Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Like I said, I'm not very knowledgeable. :-D

I just showed how daft I am, and how long it has been since I looked at
this stuff. I had looked at the image, and missed that it is on the wrong
side to be a sum. I am interested in what the real answer is. Just for
fun, I got out my old Tracor slide chart and PGT X-Ray Energy "ruler". I
poked around a bit, but nothing makes sense, unless you have Scandium in
your sample. So, I will wait for someone who knows what they are talking
about, and learn...

Regards,
Darrell



"Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu} wrote on 02/21/2013
03:44:46 PM:

} From: "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu}
} To: Darrell Miles/Fishkill/IBM-at-IBMUS,
} Date: 02/21/2013 03:47 PM
} Subject: RE: [Microscopy] Re: Subject: Unidentified extra XEDS peaks
}
} Did you check the link to the JPG file that he offered? There are
} peaks that are downscale and proportionate to the major peaks.
}
} I see sum peaks often in our Oxford system when I am looking at Al
} and push the dead time up toward 30%. I keep finding "argon" at 3 keV.
}
} -----Original Message-----
} From: milesd-at-us.ibm.com [mailto:milesd-at-us.ibm.com]
} Sent: Thursday, February 21, 2013 2:41 PM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] Re: Subject: Unidentified extra XEDS peaks
}
}
}
}
}
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}
} I am by no means very knowledgeable in the subject, but isn't that a sum

} peak? ( Ti Ka1+2)
}
} Regards,
} Darrell
}
}
} Edelmare-at-miamioh.edu wrote on 02/21/2013 11:44:39 AM:
}
} } From: Edelmare-at-miamioh.edu
} } To: Darrell Miles/Fishkill/IBM-at-IBMUS,
} } Date: 02/21/2013 11:55 AM
} } Subject: [Microscopy] Subject: Unidentified extra XEDS peaks
} }
} }
} }
} }
} }
}
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} }
} }
} } Hello listers, got an odd question that I am hoping the more
experienced
}
} } microanalytical folks can help me with.
} }
} } I think we are seeing some type of "Escape" peak using an SDD XEDS
} } system, but these are not the 1.740kev Silicon escape peaks I am used
} to.
} } They are much closer to the primary peak, vary in energy displacement
} from
} } the primary peaks, and seem to be proportionally larger than Silicon
} escape
} } peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at-
} 4.508kev),
} } 0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
} }
} } They are not "system peaks" as they track with the larger primary
peaks
} as
} } we change samples. I suspect that they are "escape artifacts" of
SDD´s
} and
} } obviously I do not spend enough time reading "MicroNews" and the
} } microanalysis literature to be aware of them. Anyone want to help us
} out,
} } please?
} }
} } We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} } 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps
(TEM
} } remember?), but have tested with same results at higher 10 - 60kcps.
} }
} } The Bruker software does not provide any identification markers. As
} they
} } are a significant size, if they are an escape artifact of SDD´s,
} } their absence
} } from the primary peak will significantly effect the quant
} } calculations. Is this
} } correct?
} }
} }
} } Thank you in advance!
} }
} }
} } 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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From: corrie.van-hoek-at-tatasteel.com
Date: Fri, 22 Feb 2013 02:39:09 -0600
Subject: [Microscopy] Subject: Unidentified extra XEDS peaks

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Dear all,

I'm 99.9% sure this is Scandium as well. The Gaussian peakshape and position matches Sc-Ka whereby the Sc-Kb is hidden under the Ti-Ka.

Some comments on the shadowing phenomena on SDD: We do have 3 SDD in house which do have shadow peaks on the left side of the main peak but this is only clearly noticable in the energy region below 1 keV. In this region, the resolution gets worse, the main peak reduces in height and a satelite peak shows up on the left side of the main peak. The peakposition of the main peak is also moving to a lower energy. Therefore analysis of B, C, N, O, and F and/or working at low kV is getting more and more problematic over time.

According to the SDD crystal supplier, X-ray photons can permanently damage the crystal structure resulting in the phenomena described. The guaranteed number of counts hitting the detector without causing damaging is 1.0E+12. This number seems to be enormous, but the reality is, using a detector at 60.000 counts/sec, 24 hrs/day, 7 days/wk, it can be used in an optimal condition for only half a year. Even just doing imaging on the microscope is contributing to the crystal damage (reported on the microscopy listserver in oct 2011).

Regards,

Corrie





________________________________________
X-from: milesd-at-us.ibm.com [milesd-at-us.ibm.com]
Sent: Thursday, February 21, 2013 23:41
To: Hoek, Corrie van

Like I said, I'm not very knowledgeable. :-D

I just showed how daft I am, and how long it has been since I looked at
this stuff. I had looked at the image, and missed that it is on the wrong
side to be a sum. I am interested in what the real answer is. Just for
fun, I got out my old Tracor slide chart and PGT X-Ray Energy "ruler". I
poked around a bit, but nothing makes sense, unless you have Scandium in
your sample. So, I will wait for someone who knows what they are talking
about, and learn...

Regards,
Darrell



"Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu} wrote on 02/21/2013
03:44:46 PM:

} From: "Straszheim, Warren E [BIOTC]" {wesaia-at-iastate.edu}
} To: Darrell Miles/Fishkill/IBM-at-IBMUS,
} Date: 02/21/2013 03:47 PM
} Subject: RE: [Microscopy] Re: Subject: Unidentified extra XEDS peaks
}
} Did you check the link to the JPG file that he offered? There are
} peaks that are downscale and proportionate to the major peaks.
}
} I see sum peaks often in our Oxford system when I am looking at Al
} and push the dead time up toward 30%. I keep finding "argon" at 3 keV.
}
} -----Original Message-----
} From: milesd-at-us.ibm.com [mailto:milesd-at-us.ibm.com]
} Sent: Thursday, February 21, 2013 2:41 PM
} To: wesaia-at-iastate.edu
} Subject: [Microscopy] Re: Subject: Unidentified extra XEDS peaks
}
}
}
}
}
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}
} I am by no means very knowledgeable in the subject, but isn't that a sum

} peak? ( Ti Ka1+2)
}
} Regards,
} Darrell
}
}
} Edelmare-at-miamioh.edu wrote on 02/21/2013 11:44:39 AM:
}
} } From: Edelmare-at-miamioh.edu
} } To: Darrell Miles/Fishkill/IBM-at-IBMUS,
} } Date: 02/21/2013 11:55 AM
} } Subject: [Microscopy] Subject: Unidentified extra XEDS peaks
} }
} }
} }
} }
} }
}
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} }
} }
} } Hello listers, got an odd question that I am hoping the more
experienced
}
} } microanalytical folks can help me with.
} }
} } I think we are seeing some type of "Escape" peak using an SDD XEDS
} } system, but these are not the 1.740kev Silicon escape peaks I am used
} to.
} } They are much closer to the primary peak, vary in energy displacement
} from
} } the primary peaks, and seem to be proportionally larger than Silicon
} escape
} } peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at-
} 4.508kev),
} } 0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
} }
} } They are not "system peaks" as they track with the larger primary
peaks
} as
} } we change samples. I suspect that they are "escape artifacts" of
SDD´s
} and
} } obviously I do not spend enough time reading "MicroNews" and the
} } microanalysis literature to be aware of them. Anyone want to help us
} out,
} } please?
} }
} } We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} } 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps
(TEM
} } remember?), but have tested with same results at higher 10 - 60kcps.
} }
} } The Bruker software does not provide any identification markers. As
} they
} } are a significant size, if they are an escape artifact of SDD´s,
} } their absence
} } from the primary peak will significantly effect the quant
} } calculations. Is this
} } correct?
} }
} }
} } Thank you in advance!
} }
} }
} } 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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28, 56 -- Subject: [Microscopy] Subject: Unidentified extra XEDS peaks
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From: johnf-at-geology.wisc.edu
Date: Fri, 22 Feb 2013 06:55:03 -0600
Subject: [Microscopy] Re: Subject: Unidentified extra XEDS peaks

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I vote for the "new artifact from new SDD" as explained by Ritchie et al
in their 2011 Microscopy and Microanalysis article (vol 17, pp 903-910):
Compton Scattering Artifacts in Electron Excited X-Ray Spectra Measured
with a Silicon Drift Detector
by Nicholas W.M. Ritchie,* Dale E. Newbury, and Abigail P. Lindstrom

(I am using it in my electron microprobe class this semester, so have a
link to it, if you need it...
{www.geology.wisc.edu/~johnf/g777/777MMarticles2.html}

John Fournelle

On 2/21/13 9:48 AM, Edelmare-at-miamioh.edu wrote:
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}
} Hello listers, got an odd question that I am hoping the more experienced
} microanalytical folks can help me with.
}
} I think we are seeing some type of "Escape" peak using an SDD XEDS
} system, but these are not the 1.740kev Silicon escape peaks I am used to.
} They are much closer to the primary peak, vary in energy displacement from
} the primary peaks, and seem to be proportionally larger than Silicon escape
} peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at- 4.508kev),
} 0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
}
} They are not "system peaks" as they track with the larger primary peaks as
} we change samples. I suspect that they are "escape artifacts" of SDD´s and
} obviously I do not spend enough time reading "MicroNews" and the
} microanalysis literature to be aware of them. Anyone want to help us out,
} please?
}
} We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps (TEM
} remember?), but have tested with same results at higher 10 - 60kcps.
}
} The Bruker software does not provide any identification markers. As they
} are a significant size, if they are an escape artifact of SDD´s, their absence
} from the primary peak will significantly effect the quant calculations. Is this
} correct?
}
}
} Thank you in advance!
}
}
} 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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--
John Fournelle
Senior Scientist
Director, Electron Probe and SEM Lab
University of Wisconsin Dept of Geoscience
Madison, Wisconsin 53706
Cell 608-438-7480


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From: philip.oshel-at-cmich.edu
Date: Fri, 22 Feb 2013 07:21:28 -0600
Subject: [Microscopy] Re: Ask-A-Microscopist regular sample with geometrical shape for

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} realname - alaa afeef
} Email - alaa.afeef-at-gmail.com
} ORGANIZATION - Glasgow University
} EDUCATION - Graduate College
} LOCATION - Glasgow
} SUBJECT_OF_QUESTION - regular sample with geometrical shape
} QUESTION -
} May I have your help with the below issue:
} I am trying to look for a sample of a 50-200 nm size to do Electron Tomography (tilted tomo),
}
} Actually, I am trying to find something that is already characterized, with geometrical shape and a well known morphology.
}
} I would be very grateful for your help.
}
} Wish you a nice weekend!
} Ala'

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From: frank_karl-at-ardl.com
Date: Fri, 22 Feb 2013 08:14:00 -0600
Subject: [Microscopy] Ask-A-Microscopist regular sample with

Contents Retrieved from Microscopy Listserver Archives
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I'd try carbon black, an ASTM N500 grade. It has the morphology and size you're interested in and depending on your definition of characterized- it's been well studied.

Failing that, I try kaolinite clay. Hexagonal shaped plates, xtal system known, crystallographic and chemical properties well documented.

My two cents..........

Stay safe.............
Frank



} May I have your help with the below issue:
} I am trying to look for a sample of a 50-200 nm size to do Electron Tomography (tilted tomo),
}
} Actually, I am trying to find something that is already characterized, with geometrical shape and a well known morphology.
}
} I would be very grateful for your help.
}
} Wish you a nice weekend!
} Ala'

==============================Original Headers==============================
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From: klivi-at-jhu.edu
Date: Fri, 22 Feb 2013 08:59:06 -0600
Subject: [Microscopy] Re: Subject: Unidentified extra XEDS peaks

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Here is a question for Richard...
When you said "Examples: 0.372 kev below Ti Ka (-at- 4.508kev), 0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev)." These were examples from different samples, each having a Ni or Cu as the major element?

Question for John Fournelle...
Can you give more specifics about the new artifact? I'm interested.
Ken

On Feb 21, 2013, at 11:47 AM, Edelmare-at-miamioh.edu wrote:

}
}
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} Hello listers, got an odd question that I am hoping the more experienced
} microanalytical folks can help me with.
}
} I think we are seeing some type of "Escape" peak using an SDD XEDS
} system, but these are not the 1.740kev Silicon escape peaks I am used to.
} They are much closer to the primary peak, vary in energy displacement from
} the primary peaks, and seem to be proportionally larger than Silicon escape
} peaks from Si\Li detectors. Examples: 0.372 kev below Ti Ka (-at- 4.508kev),
} 0.610kev below Ni Ka (-at- 7.471 kev), and 0.646 Kev Cu Ka (-at- 8.040kev).
}
} They are not "system peaks" as they track with the larger primary peaks as
} we change samples. I suspect that they are "escape artifacts" of SDD´s and
} obviously I do not spend enough time reading "MicroNews" and the
} microanalysis literature to be aware of them. Anyone want to help us out,
} please?
}
} We are working with a JEOL-2100, LaB6 at 200kV, and a Bruker Quantax
} 200 SDD. Using a beryllium holder. Count rates of 0.7 - 2.0 Kcps (TEM
} remember?), but have tested with same results at higher 10 - 60kcps.
}
} The Bruker software does not provide any identification markers. As they
} are a significant size, if they are an escape artifact of SDD´s, their absence
} from the primary peak will significantly effect the quant calculations. Is this
} correct?
}
}
} Thank you in advance!
}
}
} 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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|_|"|_|"|_|_|"|_|"|_|"|_|_|"|_|"|_|_|"|_|"|_|"|_|_|
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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From: DRK-at-shcc.org
Date: Fri, 22 Feb 2013 13:47:33 -0600
Subject: [Microscopy] Re: flat embedding large size tissue in LR White

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

To the listserver,

Some time ago was a posted query regarding embedding large tissues in LR White. As an alternative to flat embedding molds, I suggest using polyethylene Wheaton Snap Caps for LR White embedding. They are available from Fisher in 22mm diameter (06-450-201) and also in larger diameters. To drive out inherent moisture in the caps, store them in the oven prior to use. Once the sample is embedded in LR White within the caps, place these directly in a 60C oven for polymerization, without any cover, with the media directly exposed to the atmosphere within the oven. Importantly, together with the samples, place either a 500ml beaker of dry ice or a small, uncovered thermos of liquid nitrogen. The sublimation of CO2 or evaporation of nitrogen displaces enough of the atmosphere within the oven cavity for quality polymerization of the media. There may be a thin gooey layer at the top of the caps but this can be wiped clean with ethanol. The media separates easily from the caps. Including an Aclar film in the bottom of the cap works well for a crystal-clear view of the embedded sample.

Doug

Douglas R. Keene
Assistant Investigator
Micro-Imaging Center
Shriners Hospital for Children
3101 SW Sam Jackson Park Road
Portland, Oregon 97239





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From: frank_karl-at-ardl.com
Date: Fri, 22 Feb 2013 13:47:58 -0600
Subject: [Microscopy] silica sizing by TEM

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Hello Everyone,
Out of left field comes: Does anyone run an aggregate particle sizing on precipitated silica by TEM? We measure the primary particle size by TEM, but we normally have to breakup the aggregate in silica recovered from rubber by ashing. Is there an ASTM or other procedure anyone can recommend?

Thanks!!!


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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From: Jan.Ringnalda-at-fei.com
Date: Fri, 22 Feb 2013 16:25:56 -0600
Subject: [Microscopy] Re: CCD sensor size

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Hi Dmitry,

Even that nice explanation is still simplified. Nowadays, microscopes have many different modes and many different detectors, hence even a single microscope system has to be considered and many 'filters'. Depending on the mode of operation, and if it has been previously accurately aligned and calibrated, these factors will determine if the actual pixel size stored with the data is actually correct for the microscope, the kV, and the mode of operation used.

FEI has software that tries to semi-automate the calibration routine, which can be stored for any camera, any mode and any kV. However, it still requires the user to recall the file that is the correct calibration for the operational conditions that are applicable. i.e. a camera at the end of an imaging filter may be calibrated for TEM mode, however if EFTEM mode is engaged to improve the usability of this detector with a reduction in effective magnification of about 10X, the user would like the calibration to be updated automatically too.

Many camera manufacturers assume that once a calibration is done and valid for a camera this is the complete picture, furthermore usually the camera SW allows any operator to change the calibrations without verifying if 'administrator' or 'supervisor' credentials are in place, and therefore all is typically NOT well; and what is good today is not necessarily right tomorrow!

In plain language this allows any operator to 'mess' with the calibrations. Nothing can be trusted. If the camera manufacturers remain in charge of the calibrations in such a manner, it is only a matter of time before wrong calibrations are used for data analysis and interpretation which is what currently happens very frequently.

Different modes:-
LM mode
LM STEM mode
TEM mode (35mm ccd camera)
TEM mode (bottom-mount camera)
EFTEM mode (specialized projection lens series to be used in TEM mode with energy filters) Diffraction mode EFTEM Diffraction mode (specialized projection lens series to be used in diffraction mode with an energy filter) STEM mode EFTSTEM mode a small camera length series to be used for optimized STEM/EELS experiments.

And, to complete the necessary scenario's, all combinations of modes with different kV's.

Keeping systems calibrated and producing accurately calibrated images is much more complex than initially thought. Even if all calibrations are done accurately; If the sample height at the time of imaging/analysis is not correct, all of the calibrations can be undone by varying the primary focusing lens too far from the 'calibrated' values.

The FEI MagCal software goes some way towards having a properly calibrated data acquisition system by allowing a semi-automated calibration routine for all different modes and all different detectors, however users still have to be vigilant to ensure the system is correctly calibrated for the kV and mode currently selected by recalling the appropriate file. However once this file is locked in by an administrator, at least the calibrations cannot be altered within the file.

Sincerely, Jan

P.S. Disclaimer: I work for FEI, manufacturer of Electron Microscope Systems, who also develop and produce SW to aid users to correctly calibrate and quantify data obtained with electron microscope systems.


-----Original Message-----
X-from: dmitry.v.sokolov-at-gmail.com [mailto:dmitry.v.sokolov-at-gmail.com]
Sent: Tuesday, February 19, 2013 1:34 AM
To: Ringnalda, Jan

Hi Amit,

the trick with the microscopes is that they can be considered as a single "filter" between your sample and your computer screen / eye.

Calibrate the instrument against known (better if with the standard
reference) sample at the magnification and other critical parameters fixed. The subsequent calibration of the image is a breeze:
Image Size (um) = Maximum Visible Length of Reference (um) * Image Size (pixels) / Maximum Visible Length of Reference (pixels)

The pixel size at given (same as at the calibration of the instrument!) imaging conditions is calculated as:
Image Size (um) / Image Size (pixels).

Estimation of the camera pixel size:
Camera pixel size (um) = Maximum Visible Length of Reference (um) / Maximum Visible Length of Reference (pixels) / Magnification

Here we come to the need of calibration of the microscope in terms of magnification at the plane of the camera chip that may be tricky. I would rely in this case on either chip data from the manufacturer of the camera or direct measurements of the chip size under the binocular microscope. Don't forget about the LM calibration too. ;0)

This all is described at MIAWiki Knowledge Network:
http://confocal-manawatu.pbworks.com/w/page/60817289/Image%20Calibration
If something is missing or unclear, your comments/suggestions at the bottom of the page will be highly appreciated.

Please Skype me: FalconDot if I can be of more help in real time.

Cheers,
Dmitry

Advanced Knowledge Management
for MICROSCOPY and Image Analysis
________________________________
Dmitry Sokolov, Ph.D.
Mob: +64 21 063 5382
dmitry.v.sokolov-at-gmail.com

19.02.2013 18:05, amit.welcomes.u-at-gmail.com ďčřĺň:
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} -at- Warren S.
} exactly! I need to calibrate my whole TEM. From magnification to
} camera length to aberration constants. for that i need what is the
} size of my "polaroid film". since we have a ccd camera instead of
} film, I need to know the actual size of ccd sensor (will I not?)
}
} -at- rest (sorry for not addressing individually) I can caliberate at
} high mag with gold lattice image, at low mag with copper grid etc, its
} the range 80000 to 200000 which is giving bit trouble.
}
} camera: Olympus Keen View G2
}
} Gatan generally gives a caliberation sample with CCD, alas! olympus dont.
}
} With Regards
} Amit
}
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From: dmitry.v.sokolov-at-gmail.com
Date: Sat, 23 Feb 2013 04:02:48 -0600
Subject: [Microscopy] CCD sensor size

Contents Retrieved from Microscopy Listserver Archives
http://www.microscopy.com/MicroscopyListserver/MicroscopyArchives.html

Thank you Jan, that's a good point!

What you suggest below for the TEM calibration procedure is rewritten in
the Knowledge Network format and published in MIAWiki.

TEM Magnification Factors:
http://confocal-manawatu.pbworks.com/w/page/63920300/TEM%20Magnification%20Factors

TEM Modes:
http://confocal-manawatu.pbworks.com/w/page/63920396/TEM%20Modes

MagCal:
http://confocal-manawatu.pbworks.com/w/page/63920440/MagCal

TEM Calibration Best Practices:
http://confocal-manawatu.pbworks.com/w/page/63920519/TEM%20Calibration%20Best%20Practices

Your corrections, suggestions and comments would be highly appreciated.

Regarding the simplification, the degree of abstraction/details depends
on the problem to be solve. Too many details make the decision making
difficult. That is the same kind of phenomena as with the signal/noise
ratio in microscopy. I believe if users remember to have the calibration
done/applied at all the critical parameters fixed, that will do the job.
Your detailed comments however put the focus further into how namely
should the calibration be done. Many thanks for that again.

With kind regards,
Dmitry

Advanced Knowledge Management
for MICROSCOPY and Image Analysis
________________________________
Dmitry Sokolov, Ph.D.
Mob: +64 21 063 5382
dmitry.v.sokolov-at-gmail.com

23.02.2013 11:35, Jan.Ringnalda-at-fei.com ďčřĺň:
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} Hi Dmitry,
}
} Even that nice explanation is still simplified. Nowadays, microscopes have many different modes and many different detectors, hence even a single microscope system has to be considered and many 'filters'. Depending on the mode of operation, and if it has been previously accurately aligned and calibrated, these factors will determine if the actual pixel size stored with the data is actually correct for the microscope, the kV, and the mode of operation used.
}
} FEI has software that tries to semi-automate the calibration routine, which can be stored for any camera, any mode and any kV. However, it still requires the user to recall the file that is the correct calibration for the operational conditions that are applicable. i.e. a camera at the end of an imaging filter may be calibrated for TEM mode, however if EFTEM mode is engaged to improve the usability of this detector with a reduction in effective magnification of about 10X, the user would like the calibration to be updated automatically too.
}
} Many camera manufacturers assume that once a calibration is done and valid for a camera this is the complete picture, furthermore usually the camera SW allows any operator to change the calibrations without verifying if 'administrator' or 'supervisor' credentials are in place, and therefore all is typically NOT well; and what is good today is not necessarily right tomorrow!
}
} In plain language this allows any operator to 'mess' with the calibrations. Nothing can be trusted. If the camera manufacturers remain in charge of the calibrations in such a manner, it is only a matter of time before wrong calibrations are used for data analysis and interpretation which is what currently happens very frequently.
}
} Different modes:-
} LM mode
} LM STEM mode
} TEM mode (35mm ccd camera)
} TEM mode (bottom-mount camera)
} EFTEM mode (specialized projection lens series to be used in TEM mode with energy filters) Diffraction mode EFTEM Diffraction mode (specialized projection lens series to be used in diffraction mode with an energy filter) STEM mode EFTSTEM mode a small camera length series to be used for optimized STEM/EELS experiments.
}
} And, to complete the necessary scenario's, all combinations of modes with different kV's.
}
} Keeping systems calibrated and producing accurately calibrated images is much more complex than initially thought. Even if all calibrations are done accurately; If the sample height at the time of imaging/analysis is not correct, all of the calibrations can be undone by varying the primary focusing lens too far from the 'calibrated' values.
}
} The FEI MagCal software goes some way towards having a properly calibrated data acquisition system by allowing a semi-automated calibration routine for all different modes and all different detectors, however users still have to be vigilant to ensure the system is correctly calibrated for the kV and mode currently selected by recalling the appropriate file. However once this file is locked in by an administrator, at least the calibrations cannot be altered within the file.
}
} Sincerely, Jan
}
} P.S. Disclaimer: I work for FEI, manufacturer of Electron Microscope Systems, who also develop and produce SW to aid users to correctly calibrate and quantify data obtained with electron microscope systems.
}
}
} -----Original Message-----
} X-from: dmitry.v.sokolov-at-gmail.com [mailto:dmitry.v.sokolov-at-gmail.com]
} Sent: Tuesday, February 19, 2013 1:34 AM
} To: Ringnalda, Jan
} Subject: [Microscopy] Re: CCD sensor size
}
}
}
}
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} Hi Amit,
}
} the trick with the microscopes is that they can be considered as a single "filter" between your sample and your computer screen / eye.
}
} Calibrate the instrument against known (better if with the standard
} reference) sample at the magnification and other critical parameters fixed. The subsequent calibration of the image is a breeze:
} Image Size (um) = Maximum Visible Length of Reference (um) * Image Size (pixels) / Maximum Visible Length of Reference (pixels)
}
} The pixel size at given (same as at the calibration of the instrument!) imaging conditions is calculated as:
} Image Size (um) / Image Size (pixels).
}
} Estimation of the camera pixel size:
} Camera pixel size (um) = Maximum Visible Length of Reference (um) / Maximum Visible Length of Reference (pixels) / Magnification
}
} Here we come to the need of calibration of the microscope in terms of magnification at the plane of the camera chip that may be tricky. I would rely in this case on either chip data from the manufacturer of the camera or direct measurements of the chip size under the binocular microscope. Don't forget about the LM calibration too. ;0)
}
} This all is described at MIAWiki Knowledge Network:
} http://confocal-manawatu.pbworks.com/w/page/60817289/Image%20Calibration
} If something is missing or unclear, your comments/suggestions at the bottom of the page will be highly appreciated.
}
} Please Skype me: FalconDot if I can be of more help in real time.
}
} Cheers,
} Dmitry
}
} Advanced Knowledge Management
} for MICROSCOPY and Image Analysis
} ________________________________
} Dmitry Sokolov, Ph.D.
} Mob: +64 21 063 5382
} dmitry.v.sokolov-at-gmail.com
}
} 19.02.2013 18:05, amit.welcomes.u-at-gmail.com ďčřĺň:
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} }
} } -at- Warren S.
} } exactly! I need to calibrate my whole TEM. From magnification to
} } camera length to aberration constants. for that i need what is the
} } size of my "polaroid film". since we have a ccd camera instead of
} } film, I need to know the actual size of ccd sensor (will I not?)
} }
} } -at- rest (sorry for not addressing individually) I can caliberate at
} } high mag with gold lattice image, at low mag with copper grid etc, its
} } the range 80000 to 200000 which is giving bit trouble.
} }
} } camera: Olympus Keen View G2
} }
} } Gatan generally gives a caliberation sample with CCD, alas! olympus dont.
} }
} } With Regards
} } Amit
} }
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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 23 Feb 2013 19:57:51 -0600
Subject: [Microscopy] viaWWW:FEG Hitachi 4500

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Email: akomakov-at-physics.siu.edu
Name: Andrei Kolmakov

Organization: SIUC

Title-Subject: [Filtered] FEG Hitachi 4500

Message: Dear Colleague,
We are using refurbished Hitachi 4500 FEG SEM for about two years. The instrument is out of service
and I don't know when last time it was serviced. Lately new symptoms start to develop: the emission
current is constantly going down and multiple fleshing does not help –the current does not recover
or stabilize. The preset value can only be reached for a short time only after HV turned Off/ON. Any
advice. If it is FEG dying does somebody have a manual to replace, adjust it?
Thank you for your comments in advance.
Andrei

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From: microscopylistserver-noreply-at-microscopy.com
Date: Sat, 23 Feb 2013 19:59:02 -0600
Subject: [Microscopy] viaWWW:Stereomicroscope System for LEICA EM AFS2

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Email: mamiller-at-coh.org
Name: Marcia Miller

Organization: Beckman Res. Inst. City of Hope

Title-Subject: [Filtered] Stereomicroscope System for LEICA EM AFS2

Message: Dear Colleague