All, I need to purchase a closed perfusion/temperature chamber to use on a confocal/MP equipped Nikon E-800 (an upright.) I would like to be able to image cell cultures and tissues (e.g. brain slices.) After sifting through the archive and doing online searches I have come up with two, the Dvorak chamber with various temperature controllers, and the Harvard Apparatus LU-CPC-CEH Leiden chamber. Would any one like to comment on these systems, and do you have any other suggestions? Vendors (and anyone not wanting to post to the server) are welcome to contact me directly. Thanks, Tom
Thomas Moninger (thomas-moninger-at-uiowa.edu) University of Iowa Central Microscopy Research Facility (www.uiowa.edu/~cemrf) View expressed are my own.
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 1 17:01:16 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (forensic-at-mindspring.com) from on Monday, March 1, 2004 at 15:08:37 ---------------------------------------------------------------------------
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (maloneyb-at-fiu.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, March 1, 2004 at 13:40:25 ---------------------------------------------------------------------------
Email: maloneyb-at-fiu.edu Name: Barbara
Organization: FCAEM/FIU
Title-Subject: [Microscopy] [Filtered] Liposomes
Question: Dear Group - have any of you been able to successfully image any liposomes on a SEM? Thanks Barb
I do believe you could use freeze-fracture or freeze-etching with or without shadowing to visualize liposomes in modern SEMs. There are a few companies on the market who sell special attachments to SEMs, so you may process your sample (freeze, fracture, etch, shadow etc) and transfer in SEM without breaking vacuum. I did see such machine from Gatan, but I am sure other companies have similar equipment. This is a great way to visualize practically any emulsion/suspension (it's widely used in cosmetic industry). I hope it helps. Sergey No interest in any manufacturer.
At 08:20 PM 3/1/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
O.k., I'm being dense here, could someone explain to me how using a higher accellerating voltage could/would/should decrease chromatic aberations in the EM? Particluarly in the TEM. Unless we're talking really low Kev (i.e. 100ev - 1,000ev vs 100,000ev - which is why I suspect one reason why low eV in SEM's is generated by decellerating the electrons at the bottom on the lens system) why would the energy spread of the primary electron beam vary?
Thank you.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 08:09:31 2004
} } } } } } } } } } } } } } Available immediately: Electroscan E3 Environmental scanning electron } } } } } } microscope. Asking $30,000 or best offer. Buyer is responsible for } } } } } } disassembly, rigging and shipping from Ithaca, NY. The instrument is } } } } } } working, and was on a service contract until last June. There are } } } } } } numerous accessories including a peltier stage, hot-stage, 1000 lb. } } } } } } tensile stage, micromanipulator and microinjector. Two of the } } } } } } mechanical pumps are equipped with Fomblin oil as is one of the } } } } } } diffusion pumps. The Oxford X-ray detector window is broken and } } has not } } } } } } been used for several years; Reply directly to hunt-at-ccmr.cornell.edu } } } } } } or call 607-255-3789 and speak with John Hunt } } } } } } } } } } } } This item is sold where is/as is, no warranties implied or given, } } } } } } payment in full due at transfer, all packing and shipping costs are } } the buyers. } } } } } } } } } } } } John Hunt } } } } } } } } } } CCMR Microscopy Facility } } } } } 255-0108 } } }
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 08:49:14 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tomsk-at-clondiag.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 2, 2004 at 02:13:05 ---------------------------------------------------------------------------
Question: hey all, we would like to use a miniaturized confocal microscope (with fibers as connectors and pinholes) to set up an fluorescence detection apparatus. Do you know any commercial available systems which could be used...since we don`t want to invent the wheel again? Many thanks Thomas
We just covered this a few weeks ago in my course:
The equation for the diameter of the disc of least confusion (d) for chromatic aberration is:
d = Cc . alpha . (delta E / Eo)
Cc is the coefficient for chromatic aberration, alpha is the convergence angle of the beam, delta E is the energy difference, and Eo is essentially the beam energy. For thermionic emission from a tungsten filament, the initial energy of the electrons varies between about 0 and 2 eV or so -- this is, I believe, a function of variations in their intitial thermal energies within the filament.
As a result, if the accelerating voltage is 2 kV, (delta E / Eo) is 0.001; if the accelerating voltage is 20 kV, (delta E / Eo) becomes 0.0001; and if the accelerating voltage is 200 kV, (delta E / Eo) is 0.00001. Thus, the diameter of the disc of least confusion for chromatic aberration is basically inversely proportional to the accelerating voltage you're using.
Hope this helps,
Ellery
--------------- Ellery E. Frahm Research Scientist/Manager Electron Microprobe Laboratory Department of Geology & Geophysics University of Minnesota - Twin Cities Lab Website: http://probelab.geo.umn.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 09:13:26 2004
On behalf of the organizers I would like to invite you to a workshop entitled "Multidimensional data presentation techniques" which will take place on April 4 from 12:30 to 4:30 just before the opening of the "Focus on Microscopy 2004" Conference in Philadelphia.
http://www.cyto.purdue.edu/FOM2004/ - workshop web page http://www.focusonmicroscopy.org/ - FOM2004 web page
This workshop will include an interactive tutorial on the use of a variety of techniques for multidimensional microscopy data presentation. Many advanced visualization packages for microscopists are commercially available. Similarly, plenty of applications for video processing and presentations can be found on the market. However, transforming complex data sets into the actual presentation for use in lectures or in web sites is not as easy as it seems. There are a variety of tricks-of-the-trade, useful suggestions, and some very nice inexpensive or free software obtainable. We would like to share our experiences and tell you about them!
You will learn how to present static 2D images as well as 3D datasets in the most efficient way. We will show you how to produce short animations using data from confocal/MP systems in highly compressed MPEG4-based formats. You will receive a handout and CD-ROM containing key materials presented in the workshop as well as a significant number of really valuable free utility software packages.
We will demonstrate: * How to compress microscopy data. What are the pros and cons of compression? Does it affect final results? What about lossy and lossless compression? * How you should present your 3D data. How to prepare 3D image reconstruction? How to create anaglyphs? How to protect the data in an anaglyph when you compress it? How to make a movie anaglyph/animation? * How to create an animation from a 3D construction. * How to create movies that are playable in PowerPoint, on web pages, or with other media. * How to understand codecs and their associated problems. * How to edit animations/movies using high-speed command line processing. * How to you add your name, logo of your institution, or other info into the movie. * How to deal with sound overlays. * How to reproduce a movie-making process. You will learn simple command line macros that are really fast!
If you are registered for the workshop, please take some time to take our pre-workshop survey. Your participation will help us greatly with the workshop preparation. We would like to know about your expectations, your level of experience in multimedia multidimensional data presentation techniques, and the issues you consider to be important. The link to the survey is present on the workshop web page.
Bartek Rajwa
rajwa at flowcyt.cyto.purdue.edu Purdue University Cytometry Laboratories
The organizers of the workshop gratefully acknowledge the assistance of Media Cybernetics Corporation, the producer of Image-Pro Plus - an image analysis software package for fluorescence imaging, quality assurance, materials imaging, and various other scientific, medical, and industrial applications.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 09:55:27 2004
Bioptechs has a line of temperature controlled chambers, with good design for the flow so that they can handle fairly large flow volume without tearing your sample off the coverslip. They also sell temperature controlled collars for the objectives, so that your oil immersion doesn't act as a huge temperature sink.
If you are heating the objectives I strongly recommend the Boekel Warmer they carry, or its like, to keep the objectives warm at all times. You want to minimize the temperature cycling on the lenses, as it will over time induce strain on the elements. That can lead to unwanted polarization effects or other problems, very unpleasant with an expensive objective.
http://www.bioptechs.com/
Note that I'm a bit biased - I don't work for Bioptechs, but I helped test these some 15 years ago when they were first designed...
-- Kevin Ryan kevin-at-mediacy.com
-----Original Message----- } From: Moninger, Thomas [mailto:thomas-moninger-at-uiowa.edu] Sent: Monday, March 01, 2004 5:23 PM To: 'Microscopy-at-MSA.microscopy.com'
All, I need to purchase a closed perfusion/temperature chamber to use on a confocal/MP equipped Nikon E-800 (an upright.) I would like to be able to image cell cultures and tissues (e.g. brain slices.) After sifting through the archive and doing online searches I have come up with two, the Dvorak chamber with various temperature controllers, and the Harvard Apparatus LU-CPC-CEH Leiden chamber. Would any one like to comment on these systems, and do you have any other suggestions? Vendors (and anyone not wanting to post to the server) are welcome to contact me directly. Thanks, Tom
Thomas Moninger (thomas-moninger-at-uiowa.edu) University of Iowa Central Microscopy Research Facility (www.uiowa.edu/~cemrf) View expressed are my own.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 09:57:49 2004
I'm not sure if this is what you mean. But if resoln (Chromatic) = constant x focal length x semi angle x delta V/V then if a finite change in voltage occurs (delta V) increasing V must improve resolution. Part of this may due to the optics of the system but a major consideration is the thickness of specimen where I'd always understood that the loss of voltage was roughly finite (I won't say linear) then increasing the overall accelerating voltage should improve resolution. This does work even at 60kv+.
I hope I haven't baffled anyone with my 'Noddy' science.
Malcolm
Malcolm Haswell e.m. unit School of Health, Natural and Social Sciences Fleming Building University of Sunderland Tyne & Wear UK
----- Original Message ----- } From: Richard Edelmann {edelmare-at-muohio.edu}
I am thinking that the energy spread of the the source should be the same reguardless of accellerating voltage and the accellerator adds the exact same elctron volts to all electrons so the energy (chromatic) spread arriving at any lens should be the same no matter the accelerating voltage. Chromatic aberation, however, is a function of the relative energy spread compared to the final ev. The deflection error due to the spread at the source has a much smaller contribution to the total deflection in a higher voltage TEM lens.
My 2 ev worth
Dave
-- David Barnard Wadsworth Ctr NYS Dept Health Albany NY 12201-0509 barnard-at-wadsworth.org 518 473-5299 voice 518 474-7992 fax
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 12:12:03 2004
Here is the table of contents for the March/April 2004 issue of Microscopy Today.
New Subscriptions via http://www.microscopy-today.com only, please. New subscriptions will close on Friday March 5th for this issue.
WE ARE PURGING NON-MSA MEMBER, NON-NORTH AMERICAN, UNSUBSCRIBED INDIVIDUALS! WE ARE AWAITING THE LATEST INPUT FROM THE MSA MEMBER RENEWAL PROCESS BEFORE COMPLETING THE PURGING PROCESS. SEVERAL HUNDRED NON-QUALIFIED SUBSCRIBERS HAVE ALREADY BEEN DROPPED.
March/April Microscopy Today Contents:
Carmichael and Lingle: Fluorescent Speckle Microscopy
Hall: The Nematode Caenorhabditis elegans, A Model Animal "Made for Microscopy"
Sedgewick: Digital Movies for Others to See
Galvez, Giberson, & Cardiff: Microwave Mechanisms - The Energy/Heat Dichotomy
Neal: Photoshop and 12-bit Digital Microscope Camera Images
Nester: Guidance for Choosing When to Use Electron and/or Light Microscopy and Related ASTM E4 Standards
Simmons: The Role of Microscopy in Indoor Air Quality Investigations
Humphrey: How to Promote a Facility in Order to Increase Use, Acquire New Equipment and, as a Result, Increase Revenue
Hudson, Benedict, & Flaitz: TEM Specimen Preparation Technique for Small Semiconductor Devices
Duke: Lens Cleaning - Best Practices Review
Stephenson and Gabel: Use of Fishing Weight Putty for Quickly Mounting SEM Specimens
Sepsenwol: A Homemade Vacuum Forceps For Mounting Small SEM Samples
Ron Anderson, Editor Microscopy Today
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 13:27:10 2004
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O.k., I'm being dense here, could someone explain to me how using a higher accellerating voltage could/would/should decrease chromatic aberations in the EM? Particluarly in the TEM. Unless we're talking really low Kev (i.e. 100ev - 1,000ev vs 100,000ev - which is why I suspect one reason why low eV in SEM's is generated by decellerating the electrons at the bottom on the lens system) why would the energy spread of the primary electron beam vary?
Thank you.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 13:47:04 2004
------------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} } } } } } } } } } } } } } Available immediately: Electroscan E3 Environmental scanning electron } } } } } } microscope. Asking $30,000 or best offer. Buyer is responsible for } } } } } } disassembly, rigging and shipping from Ithaca, NY. The instrument is } } } } } } working, and was on a service contract until last June. There are } } } } } } numerous accessories including a peltier stage, hot-stage, 1000 lb. } } } } } } tensile stage, micromanipulator and microinjector. Two of the } } } } } } mechanical pumps are equipped with Fomblin oil as is one of the } } } } } } diffusion pumps. The Oxford X-ray detector window is broken and } } has not } } } } } } been used for several years; Reply directly to hunt-at-ccmr.cornell.edu } } } } } } or call 607-255-3789 and speak with John Hunt } } } } } } } } } } } } This item is sold where is/as is, no warranties implied or given, } } } } } } payment in full due at transfer, all packing and shipping costs are } } the buyers. } } } } } } } } } } } } John Hunt } } } } } } } } } } CCMR Microscopy Facility } } } } } 255-0108 } } }
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 13:50:22 2004
I am the keeper of the MSA videos and am in the process of transferring them to DVD. Several masters are in bad condition and I do not have any other copies. If there is anyone out there who has a copy of number 214, 3D Deconvolution from 1998, I would appreciate the opportunity to borrow it so that it may be restored to the collection. You may expect to hear from me in the future with similar requests. Thanks in advance
Greg Erdos
Gregory W. Erdos Ph.D. Assistant Director, Biotechnology Program Scientific Director, Electron Microscopy P.O. Box 118525 217 Carr Hall University of Florida Gainesville, FL 32611 gwe-at-ufl.edu 352-392-1295 fax- 352-846-0251
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 14:11:32 2004
Try contacting Martin Harris at Optiscan Imaging, Melbourne, Australia (martinh-at-optiscan.com) they specialise in miniture confocal instruments for edoscopy and the like.
Ian
Ian Hallett HortResearch Mt Albert Research Centre, Private Bag 92 169 Auckland, New Zealand Fax +64 9 815 4201 Telephone +64 9 815 4200 EMail ihallett-at-hortresearch.co.nz
} } } by way of MicroscopyListserver {tomsk-at-clondiag.com} 3/03/2004 4:02:27 } } }
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (tomsk-at-clondiag.com) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 2, 2004 at 02:13:05 ---------------------------------------------------------------------------
Question: hey all, we would like to use a miniaturized confocal microscope (with fibers as connectors and pinholes) to set up an fluorescence detection apparatus. Do you know any commercial available systems which could be used...since we don`t want to invent the wheel again? Many thanks Thomas
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From MicroscopyL-request-at-ns.microscopy.com Tue Mar 2 17:23:06 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (njws-at-aol.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 2, 2004 at 15:56:37 ---------------------------------------------------------------------------
Question: For anyone in the Baltimore/Washington area or anyone else willing to pick them up, I have the following items to donate free of charge. I have retired after 40 years in the field of electron microscopy and would like not to have to leave these things at the curb for regular Thursday pick-up. All must be taken(not piecemeal) and no shipping. Anyone interested should e-mail me to set up a time.
BOOKS: Biology of animal viruses,vol I, II Histological techniques for electron microscopy A color atlas and text for histopathology Immunology for students of medicine Electron microscopic anatomy Positive staining for electron microscopy X-ray microanalysis in the electron microscope Principles and techniques of electron microscopy,Hyatt Principles and techniques in histochemistry Ultramicrotomy,Reid Cell pathology Ultrastructural pathology of the cell An introduction to virology Diagnostic electron microscopy Viral immunodiagnosis Low temperature methods in electron microscopy Histology-a text and atlas,Rhodin A textbook of histology,Bloom and Fawcett An atlas of ultrastructure,Rhodin Ultrastructural aspects of disease
OTHER:
Ohaus triple bneam balance Fisher water bath 2 stereo heads approx8 boxes various sized glass strips (LKB)
There was a virus attack on Dee Breger's computer.
The virus is sending out alot of email with her name on it and since the faked Email address is one of a valid "subscriber" the messages are getting through the filter.
There was a slew of them which appeared between ~ 5:30-6 PM Central Standard Time on 3/2/04
Please trash any Email with her name on it immediately. I am in the process of trying to block any new postings.
Please be aware that spam attacks like this frequently are intended to embarrass the person whose "ID" was stolen/forged. Or to use them as a source of continued infection. Do not reply, I would even suggest that you do not even open the messages.
On behalf of Dee, who has also called me, she apologizes that this has happened and hopes that you will all understand.
Nestor Your Friendly Neighborhood SysOp
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 02:58:05 2004
-- Hi everybody, We have to confirm the identity of polypropylene and polyethylene in a film embedded in a resin using TEM. We cut with the ultramicrotome the sections and stain with OsO4 during 30 minutes. It is the first time that we do this and the results aren´t good, because there isn´t reaction with the OsO4. Has everyone done this type of work? We need sugerences. A lot of thanks in advance
************************************************************* Dra. María Belén López Mosquera Unidade de Microscopía Servicios Xerais de Apoio á Investigación Universidade da Coruña Edificio Servicios Centrais de Investigación Campus de Elviña s/n E-15071 A Coruña
} From 'Polymer Microscopy' by Saywer and Grubb I got the idea that it's not the energy spread of the primary beam that limits resolution but the energy spread due to the specimen. I assume that the same formula for the disk of least confusion applies no matter what the source of deltaE, but I'm wondering which energy spread is really limiting for high resolution TEM of 2D protein crystals for example.
Here comes a somewhat lengthy discussion that I wrote when I read that book. I'd be grateful if somebody could tell me whether I'm on the right track:
The most probable inelastic interaction is the 'plasmon production' with a most probable energy
loss to the scattered electrons of around 25 eV for carbon (EELS Atlas).
The mean free path for plasmon loss scattering is in the order of 100 nm for 100 keV electrons
(table in Williams and Carter, page 657, which unfortunately doesn't include carbon).
I think the argument used in 'Polymer Microscopy' by Sawyer and Grubb runs as follows:
} From the data above an electron loses about 25 eV of energy every 100 nm or 0,25 eV per nm of
specimen thickness it traverses.
Since this energy loss is stochastic the beam also suffers an energy-spread deltaE of about
the same magnitude. Due to the chromatic aberration of the objective lens this energy-spread limits the
attainable resolution to a value that depends on the thickness given by the following rule of thumb:
One should not expect to resolve details smaller than one tenth of the specimen thickness for
carbonacious specimens.
My comment: This argument is only valid for specimens that are at least 100 nm thick. If the specimen
is considerably thinner than the mean free path for inelastic scattering (for example 10-20 nm as is usual
in structure determination of proteins (excluding viruses)) only relatively few electrons have been
inelastically scattered at all and it doesn't make sense to speak of an energy spread due to the specimen.
The unscattered (which form the majority) and elastically scattered electrons have the same energy-spread
as the incoming electrons. Most of the inelastically scattered electrons have an energy spread that's given
by the width of the plasmon loss curve, that means a deltaE of about 50 eV (Ahn, Krinanek: EELS-Atlas).
In the weak phase-weak amplitude approximation only the unscattered and elastically scattered electrons
are regarded as taking part in image formation, whereas the inelastically scattered electrons lead to
background noise. Therefore the energy-spread of the inelastically scattered electrons is irrelevant.
Sawyer and Grubb treat energy-loss as a continuous process (". will cause a 100 keV electron to loose
about 0.25 eV per nanometer of thickness on average."). In fact an electron either losses the energy of a
plasmon (10 to 50 eV) or it doesn't and on average it does so every 100 nm it travels through the specimen.
Only when the specimen is thicker than the mean free path does it make sense to speak of an average
energy loss per nm thickness.
For faster electrons (for example 300 keV) the mean free paths are longer and accordingly all thickness
limits higher.
Philip Koeck Svdertvrns Hvgskola and Karolinska Institutet Dept. of Bioscience at Novum S-14157 Huddinge Sweden phone: +46-8-6089186 fax: +46-8-6089290 http://www.biosci.ki.se/em
'This winter I am giving courses to three students, of whom one is only moderately prepared, the other less than moderately, and the third lacks both preparation and ability. Such are the burdens ...' Carl Friedrich Gauss (1810) ______________________________________________
----- Original Message ----- } From: "Ellery Frahm" {frah0010-at-umn.edu} To: {edelmare-at-MUOHIO.EDU} ; {microscopy-at-MSA.Microscopy.com} Sent: Tuesday, March 02, 2004 4:07 PM
Folks:
Thank you for the info! I was not aware of the chromatic aberation equation. Thank you.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 08:20:42 2004
Hello: I would try staining with Ruthenium Tetraoxide (RuO4). Even though both polymers will stain they should stain at different rates. According to "Ruthenium Tetraoxide Staining of Polymers for Electron Microscopy" by Trent et al published in Macromolecules 1983 16, 589-598 polyethylene oxide will stain faster than isotactic polypropylene. I would stain only for between 5 min and 15 min. Both RuO4 and OsO4 are very dangerous chemicals so please be sure to observe proper safety procedures. Also you will need to use a cryo-ultramicrotome or the polypropylene phase will smear. Good luck. Steve
-- Hi everybody, We have to confirm the identity of polypropylene and polyethylene in a film embedded in a resin using TEM. We cut with the ultramicrotome the sections and stain with OsO4 during 30 minutes. It is the first time that we do this and the results aren´t good, because there isn´t reaction with the OsO4. Has everyone done this type of work? We need sugerences. A lot of thanks in advance
Stephen McCartney Research Associate Materials Research Institute 2108 Hahn Hall Va Tech Blacksburg, VA 24061 540-231-9765 - phone 540-231-8517 - FAX
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 08:44:22 2004
To first-order plasmons don't change the angle of the electrons, so an image with the plasmons is very close to an image with the no-loss electrons shifted in focus (the chromatic aberration term). A rough approximation is to consider the electrons which have lost energy as a background term, say B, in which case your contrast goes down by ~1/(1+B) so you lose apparent resolution because your signal-to-noise ratio is getting worse. (Noise from counting statistics; there is only one electron in the column at a time and you don't collect that many in an image.) For a thicker sample you should also reduce the amplitude/intensity of the zero- loss electrons, another reduction in your contrast. Note that it is contrast relative to noise that matters. (There are some non-linear imaging effects, and the imaging theory for electrons which have lost energy is a bit more complicated than this, but these effects is probably not relevant for proteins.)
N.B. I suspect that you are understimating the mean-free path, particularly for a low-density protein.
----------------------------------------------- Laurence Marks Department of Materials Science and Engineering Northwestern University Evanston, IL 60201, USA Tel: (847) 491-3996 Fax: (847) 491-7820 mailto:ldm2-at-risc4.numis.nwu.edu http://www.numis.nwu.edu -----------------------------------------------
On Wed, 3 Mar 2004, Philip Koeck wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } A related question: } } } From 'Polymer Microscopy' by Saywer and Grubb I got the idea that it's not } the energy } spread of the primary beam that limits resolution but the energy spread due } to the specimen. } I assume that the same formula for the disk of least confusion applies no } matter what the source } of deltaE, but I'm wondering which energy spread is really limiting for high } resolution TEM } of 2D protein crystals for example. } } Here comes a somewhat lengthy discussion that I wrote when I read that book. } I'd be grateful } if somebody could tell me whether I'm on the right track: } } The most probable inelastic interaction is the 'plasmon production' with a } most probable energy } } loss to the scattered electrons of around 25 eV for carbon (EELS Atlas). } } The mean free path for plasmon loss scattering is in the order of 100 nm for } 100 keV electrons } } (table in Williams and Carter, page 657, which unfortunately doesn't include } carbon). } } } } I think the argument used in 'Polymer Microscopy' by Sawyer and Grubb runs } as follows: } } } From the data above an electron loses about 25 eV of energy every 100 nm or } 0,25 eV per nm of } } specimen thickness it traverses. } } Since this energy loss is stochastic the beam also suffers an energy-spread } deltaE of about } } the same magnitude. Due to the chromatic aberration of the objective lens } this energy-spread limits the } } attainable resolution to a value that depends on the thickness given by the } following rule of thumb: } } One should not expect to resolve details smaller than one tenth of the } specimen thickness for } } carbonacious specimens. } } } } My comment: This argument is only valid for specimens that are at least 100 } nm thick. If the specimen } } is considerably thinner than the mean free path for inelastic scattering } (for example 10-20 nm as is usual } } in structure determination of proteins (excluding viruses)) only relatively } few electrons have been } } inelastically scattered at all and it doesn't make sense to speak of an } energy spread due to the specimen. } } The unscattered (which form the majority) and elastically scattered } electrons have the same energy-spread } } as the incoming electrons. Most of the inelastically scattered electrons } have an energy spread that's given } } by the width of the plasmon loss curve, that means a deltaE of about 50 eV } (Ahn, Krinanek: EELS-Atlas). } } In the weak phase-weak amplitude approximation only the unscattered and } elastically scattered electrons } } are regarded as taking part in image formation, whereas the inelastically } scattered electrons lead to } } background noise. Therefore the energy-spread of the inelastically scattered } electrons is irrelevant. } } Sawyer and Grubb treat energy-loss as a continuous process (". will cause a } 100 keV electron to loose } } about 0.25 eV per nanometer of thickness on average."). In fact an electron } either losses the energy of a } } plasmon (10 to 50 eV) or it doesn't and on average it does so every 100 nm } it travels through the specimen. } } Only when the specimen is thicker than the mean free path does it make sense } to speak of an average } } energy loss per nm thickness. } } } } For faster electrons (for example 300 keV) the mean free paths are longer } and accordingly all thickness } } limits higher. } } } } } } } } Philip Koeck } Svdertvrns Hvgskola and } Karolinska Institutet } Dept. of Bioscience at Novum } S-14157 Huddinge } Sweden } phone: +46-8-6089186 } fax: +46-8-6089290 } http://www.biosci.ki.se/em } } 'This winter I am giving courses to three students, of whom one is only } moderately prepared, } the other less than moderately, and the third lacks both preparation and } ability. } Such are the burdens ...' Carl Friedrich Gauss (1810) } ______________________________________________ } } } ----- Original Message ----- } } From: "Ellery Frahm" {frah0010-at-umn.edu} } To: {edelmare-at-MUOHIO.EDU} ; {microscopy-at-MSA.Microscopy.com} } Sent: Tuesday, March 02, 2004 4:07 PM } Subject: [Microscopy] Re: KeV vs chromatic aberation } } } } } } } } -------------------------------------------------------------------------- } ---- } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -------------------------------------------------------------------------- } ----- } } } } Hi Richard, } } } } We just covered this a few weeks ago in my course: } } } } The equation for the diameter of the disc of least confusion (d) for } } chromatic aberration is: } } } } d = Cc . alpha . (delta E / Eo) } } } } Cc is the coefficient for chromatic aberration, alpha is the convergence } } angle of the beam, delta E is the energy difference, and Eo is essentially } } the beam energy. For thermionic emission from a tungsten filament, the } } initial energy of the electrons varies between about 0 and 2 eV or so -- } } this is, I believe, a function of variations in their intitial thermal } } energies within the filament. } } } } As a result, if the accelerating voltage is 2 kV, (delta E / Eo) is 0.001; } } if the accelerating voltage is 20 kV, (delta E / Eo) becomes 0.0001; and } if } } the accelerating voltage is 200 kV, (delta E / Eo) is 0.00001. Thus, the } } diameter of the disc of least confusion for chromatic aberration is } } basically inversely proportional to the accelerating voltage you're using. } } } } Hope this helps, } } } } Ellery } } } } --------------- } } Ellery E. Frahm } } Research Scientist/Manager } } Electron Microprobe Laboratory } } Department of Geology & Geophysics } } University of Minnesota - Twin Cities } } Lab Website: http://probelab.geo.umn.edu } } } } } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 09:20:03 2004
We are looking for a new critical point dryer and would appreciate any feedback users can provide. We currently have a Polaron Jumbo, that is about 30 years old. It works, but the temperature rises very high - it is controlled just by hot water - and each run seems to have different conditions. We would like to get a machine that might be more constant from one run to the next, and that students might find somewhat easier to use. Most of the CPDs we have seen through our web hunts seem to have very small sample chambers, though. We would like to know 1) what CPDs people like, 2) whether sample chamber size seems to be a problem, 3) any other concerns that users have and would alert us to. Many thanks,
Dr. Sally Leys Assistant Professor and Canada Research Chair in Evolutionary Developmental Biology Department of Biological Sciences CW 405 The University of Alberta Edmonton, Alberta Canada T6G 2E9
How can we best prepare our soil samples for SEM and micro analysis? We are novices at this. We have great equipment (Hitachi FESEM 4500 and EDAX Genesis) We are interested in phosphorus and iron in river sediments. We are having difficulty in interpreting the results we have been getting (we crushed soil to double sticky tape, mounted on Al stubs and coated with AuPd). The preps look good at the SEM with little charging.
What we see is grains and some organic matter (OM) and coatings. The spectrum shows some OM and SiO2 along with other particles of mixed composition.
My question is how can we interpret this? What about references etc? We need your expertise!
Your help will be greatly appreciated. Carol Bronick
Agricultural Research Station Box 9061 Virginia State University Petersburg, VA 23806 804.524.6822 cbronick-at-vsu.edu
____________________________________________________________________ Check your SchoolEmail at http://www.CampusI.com Find the LOWEST PRICES on books at http://www.campusi.com/BookFind !
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 09:57:51 2004
Slight correction to my previous post. Of course you will have problems with both the polyethylene and polypropylene if you don't use a cryo system when microtoming. Steve
Hello: I would try staining with Ruthenium Tetraoxide (RuO4). Even though both polymers will stain they should stain at different rates. According to "Ruthenium Tetraoxide Staining of Polymers for Electron Microscopy" by Trent et al published in Macromolecules 1983 16, 589-598 polyethylene oxide will stain faster than isotactic polypropylene. I would stain only for between 5 min and 15 min. Both RuO4 and OsO4 are very dangerous chemicals so please be sure to observe proper safety procedures. Also you will need to use a cryo-ultramicrotome or the polypropylene phase will smear. Good luck. Steve
Stephen McCartney Research Associate Materials Research Institute 2108 Hahn Hall Va Tech Blacksburg, VA 24061 540-231-9765 - phone 540-231-8517 - FAX
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 10:52:52 2004
Dear Sally, I recently needed to buy a CPD for my new lab and after looking into the options I bought a Tousimis Samdri PVT-3D. This unit has cooling for the chamber provided by letting CO2 from the tank run past the chamber, so it is easy to cool. I like not having to mess with water cooling. I also like to do my ethanol CO2 exchanges around 0C. The chamber size is 1.25 in round by 1.25 in tall. There is an accessory available to make a much bigger chamber (not wider, just longer). I also have been extremely impressed by the Tousimis company support, with nice instructions and extras, and prompt responses to my frequent questions.
I have no financial interest in the company, just a happy customer,
Tobias
} } We are looking for a new critical point dryer and would appreciate } any feedback users can provide. We currently have a Polaron Jumbo, } that is about 30 years old. It works, but the temperature rises very } high - it is controlled just by hot water - and each run seems to } have different conditions. We would like to get a machine that might } be more constant from one run to the next, and that students might } find somewhat easier to use. Most of the CPDs we have seen through } our web hunts seem to have very small sample chambers, though. We } would like to know 1) what CPDs people like, 2) whether sample } chamber size seems to be a problem, 3) any other concerns that users } have and would alert us to. } Many thanks, } } Dr. Sally Leys } Assistant Professor and Canada Research Chair } in Evolutionary Developmental Biology } Department of Biological Sciences CW 405 } The University of Alberta } Edmonton, Alberta } Canada } T6G 2E9 } } Telephone: (780) 492-6629 } Fax: (780) 492-9234 } Email: sleys-at-ualberta.ca
The previous responders have made mostly made good points, but have neglected one important possibility. Zeiss planapo lenses of that era are of high optical quality, but unfortunately they frequently suffer from a separation of lens elements known as delamination, caused by a failure of the lens cement. Used objectives also frequently have dust or a deposit of fine, misty oil droplets on the surface of the back lens. If you have a "phase telescope" you should focus through the objective and look for contamination, or delamination, which may appear as gray patches or show interference colors, and usually start at the edges and work their way inward. You can also use a dissecting microscope to look through the objective (put a light source in front of the objective and look through the back).
Either contamination or delamination will cause a haziness of the image, which will be reduced with stopping down of the condenser, explaining why the best image is seen when the condenser is stopped down past it's theoretical optimal position. Stopping down also causes dirt or other artifacts in the optical path to become much more noticeable. If you can't locate the points of contamination with the phase telescope, try rotating the objective to see if the artifacts move with it. Haze, dirt, and delamination may also be present in the condenser. Most used objectives and condensers I have come across were in need of a good cleaning.
If you want to use your 100x planapo or any other oil lens, not using oil between the specimen and the objective is not an option. With my Zeiss microscope and 1.4 NA condenser, oiling the condenser noticeable improves the contrast as well as resolution. As others have said, you should definitely not oil your 0.9 NA condenser. Also be aware that different types of immersion oil should never be allowed to mix, and that some types of oil can damage the mounting cement of some brands of lenses. Using Zeiss immersion oil with some Nikon lenses, for example, can be disastrous.
Ralph Common Electron Microscopist Michigan State University Division of Human Pathology A608 East Fee Hall East Lansing, MI 48824 517-355-7558; fax 517-432-1053 ralph.common-at-ht.msu.edu
-----Original Message----- } From: Bruce Girrell [mailto:bigirrell-at-microlinetc.com] Sent: Thursday, February 26, 2004 5:08 PM To: Microscopy-at-msa.microscopy.com
I have a binocular Zeiss phase contrast microscope that came with a basic assortment of lenses. A while back I bought a planapo 10/0.32 lens (standard - no phase ring) and was stunned at the quality of the image. To describe the image as "sharp" or even "crisp" doesn't do it justice. Some stained thin sections have the appearance of stained glass and the image almost takes on a three dimensional quality. I was impressed.
Recently, I was able to acquire a planapo 100/1.3 Ph3. Despite its very impressive appearance, I have been much less impressed with the image produced by this lens. Granted, the contrast is better than the standard lens, but based on the difference I had seen at 10x I was expecting a little more in the sharpness department.
Bear in mind that I fall in the "hobbyist" ranks. Some of these questions are basic. I have set up Kohler illumination. I believe that I have the phase rings aligned. Most of these questions apply to brightfield observation anyway.
Questions: 1) Am I simply expecting too much at 1000x? 2) Is the phase ring degrading the performance? 3) My condenser only has an n.a. of 0.9. Is this the problem? 4) Even with this condenser, should I be using oil on it as well as the objective? 5) With the phase stop in place performance is worse - lots of artifacts. That is not the case with the 40x and 10x phase objectives.
One more general question: Literature tells me that I should achieve optimum contrast/resolution with the brightfield aperture closed about 1/3 of the way. I seem to get the best results with the aperture closed just a little over half way. Am I not sensitive enough to artifacts and confusing increased contrast for increased resolution? Am I maybe using too much light?
Thanks for your help,
Bruce Girrell
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 13:23:26 2004
This is a question about TEM objective apertures, mostly about thickness and selection of hole size.
We have a JEOL 1200, I wanted to replace the objective aperture, old one was not cleaning up well. Ordered one from EM supplier we get most everything else from. Picked one that was for JEOL, a Pt strip .1 mm thick, three holes, 50, 100, 150. Sounded perfect.
Tried to put it in yesterday. Way too thick. Aperture holder accepts the strip by slipping it in a fixed sandwich kind of space, not with a separate screw down holder over the top. Old aperture was way thinner than the new one, even though new one was called 'thin'.
Checked all the other catalogs I have, most had apertures with the same dimensions. Don't have much experience with this so I am hoping to be educated.
Are the apertures for this instrument something special? Where do I get an aperture to fit? Is JEOL the only source? What criteria do I use to select the hole size? etc.
Help me out on this one.
Thanks
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 13:38:11 2004
First of all, Au lines will overlap with P, so in this case coating with Au is not acceptable.
Then I am not sure about a goal of your research. If you need just to determine whether Fe and P are in your samples, then EDS is not the best choice of method. Depending on anticipated levels of elements the XRF, wet chemistry or even mass spectrometry could be a better choice.
If detection limit of 0.5% is OK, then I would use high intensity beam current (so that dead time is about 20-50%)to acquire spectrum from pretty big field of view, at magnification of x100 or x200, for at least 5 min. Sometimes fast mapping (again at high beam current and at magnifications when many particles are visible) can help to localize the place of interest.
Regards,
Vladimir
Vladimir M. Dusevich, Ph.D. Electron Microscope Lab Manager 3127 School of Dentistry 650 E. 25th Street Kansas City, MO 64108-2784
} Hello fellow Microscopists, } } How can we best prepare our soil samples for SEM and micro } analysis? We are novices at this. We have great equipment } (Hitachi FESEM 4500 and EDAX Genesis) We are interested in } phosphorus and iron in river sediments. We are having } difficulty in interpreting the results we have been getting } (we crushed soil to double sticky tape, mounted on Al stubs } and coated with AuPd). The preps look good at the SEM with } little charging. } } What we see is grains and some organic matter (OM) and } coatings. The spectrum shows some OM and SiO2 along with } other particles of mixed composition. } } My question is how can we interpret this? What about } references etc? We need your expertise! } } Your help will be greatly appreciated. } Carol Bronick } } Agricultural Research Station } Box 9061 } Virginia State University } Petersburg, VA 23806 } 804.524.6822 } cbronick-at-vsu.edu } } ____________________________________________________________________ } Check your SchoolEmail at http://www.CampusI.com } Find the LOWEST PRICES on books at http://www.campusi.com/BookFind ! } }
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 14:05:24 2004
Why don't you use JEOL spares as your first choice?
At least you then know that they will fit and work.
Down here, at least, JEOL parts are not exhorbitantly expensive.
cheers
rtch (a satisfied customer of JEOL Sydney)
Date sent: Wed, 3 Mar 2004 11:17:01 -0800 To: Microscopy-at-sparc5.microscopy.com } From: jmkrupp-at-cats.ucsc.edu (Jon Krupp)
Jonathan I do believe, the apertures in JEM1200 are made from molybdenum (or tungsten?), definitely not a Pt. Perhaps you have to contact JEOL for specification (they are quite friendly). Personally, I would avoid "after market" spare parts especially objective apertures (they are designed in the way they are actually self-cleaning under the beam). Aperture in my microscope is "in" for more than 5 years and I never cleaned it. Deposits on the objective aperture usually indicates problem with vacuum. You may easily clean original JEOL aperture by heating up in tantalum or tungsten boat in good vacuum (5x10-6 torr or better) for a minute or less. It cleans everything. The problem there is that it's possible to damage the hole itself (well, it means, it's time to buy new one) if overheat. It's important, the boat made from different material than aperture, otherwise, the strip will stick to the boat. I am using 25-50-100 um objective aperture strip. In 95% cases we are using 50um spot - it provides enough light with decent contrast. I am using 25um spot (not recommended to use with plastic sections!) for very weak W shadowed samples only . I never used 100 um. I hope it helps. Sergey
At 11:17 AM 3/3/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
Actually, P K alpha and Au M alpha don't overlap but are indeed very close by 108eV. Pt might be an option but its M alpha is even closer to the P K alpha. The 108eV distance at low eV is not always a problem with Genesis. The EDAX HPD feature is very powerful for deconvoluting peaks. A quant with low intensity errors will give very good results. Any element that has high (} 20%) intensity error should be discarded or re-run at different KV.
At F, my detector is about 56-59eV (give or take) resolution and how one calculates this. Calibrated rez at Al and Cu is typically 128eV at 102uS. No spec wars please.
Potting the soil in metallurgical mounting media then polishing should provide good results. The irregularity of un-polished (3-D) soil grains is not good for EDS. Flat, polished surfaces are best.
Collect at 102uS with at least 1,000cps and do it for about 300 live seconds. Keep DT { 25%. Then do peak ID, HPD and finally, quant and look at the Inten. Error values.
I look at C, F, O, P, Al, Si, Fe, S, Cu, W, Ta, Hf, Zr, and many others (not all at the same time!!) that are Au/Pd coated. At 2X KV of highest value, I can't recall a time that I could not define the constituents with EDAX EDS. Organics/polymers are another story-- need FTIR or WDS for that.
gary g.
At 11:51 AM 3/3/2004, Dusevich, Vladimir wrote:
} First of all, Au lines will overlap with P, so in this } case coating with Au is not acceptable. } } Then I am not sure about a goal of your research. } If you need just to determine whether Fe and P are in } your samples, then EDS is not the best choice of method. } Depending on anticipated levels of elements the XRF, } wet chemistry or even mass spectrometry could be a better } choice. } } If detection limit of 0.5% is OK, then I would use } high intensity beam current (so that dead time is about } 20-50%)to acquire spectrum from pretty big field of view, } at magnification of x100 or x200, for at least 5 min. } Sometimes fast mapping (again at high beam current and at } magnifications when many particles are visible) can } help to localize the place of interest. } } Regards, } } Vladimir } } } Vladimir M. Dusevich, Ph.D. } Electron Microscope Lab Manager } 3127 School of Dentistry } 650 E. 25th Street } Kansas City, MO 64108-2784 } } Phone: (816) 235-2072 } Fax: (816) 235-5524 } Web: http://www.umkc.edu/dentistry/microscopy } } } } Hello fellow Microscopists, } } } } How can we best prepare our soil samples for SEM and micro } } analysis? We are novices at this. We have great equipment } } (Hitachi FESEM 4500 and EDAX Genesis) We are interested in } } phosphorus and iron in river sediments. We are having } } difficulty in interpreting the results we have been getting } } (we crushed soil to double sticky tape, mounted on Al stubs } } and coated with AuPd). The preps look good at the SEM with } } little charging. } } } } What we see is grains and some organic matter (OM) and } } coatings. The spectrum shows some OM and SiO2 along with } } other particles of mixed composition. } } } } My question is how can we interpret this? What about } } references etc? We need your expertise! } } } } Your help will be greatly appreciated. } } Carol Bronick } } } } Agricultural Research Station } } Box 9061 } } Virginia State University } } Petersburg, VA 23806 } } 804.524.6822 } } cbronick-at-vsu.edu } } } } ____________________________________________________________________ } } Check your SchoolEmail at http://www.CampusI.com } } Find the LOWEST PRICES on books at http://www.campusi.com/BookFind ! } } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 3 15:39:14 2004
In response to Sergey and Jonathan's questions let me say this: Ladd, as most people know, produces probably 97% of all apertures made in the U.S. We customize apertures for all microscopes to fit the user's needs. For instance in Sergey's case, he could have two or even three 50um holes if he wanted. The "after market" apertures offer you this customization opportunity. We can machine in various materials, such as moly, platinum, etc. and provide strips or discs as thin as 0.025mm.
Generally, apertures in the "after market" are quite a bit less expensive, more readily available and, as I mentioned, can be customized to meet your needs. We put as many as 20 holes in some plates and strips for specialized equipment.
As a matter of fact most apertures are "after market" because EM manufacturers tend not to produce their own apertures. I am not impartial on this matter, but for flexibility and value the "after market" is the place to look for apertures.
Disclaimer: Ladd Research sells EM supplies including apertures and specialized micro-holes to EM users and microscope manufacturers.
John Arnott
Ladd Research 83 Holly Court Williston, VT 05495
On-line Catalog: http://www.laddresearch.com
tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) fax: 1-802-660-8859 e-mail: sales-at-laddresearch.com ----- Original Message ----- } From: "Sergey Ryazantsev" {sryazant-at-ucla.edu} To: {Microscopy-at-sparc5.microscopy.com} Sent: Wednesday, March 03, 2004 3:56 PM
John I LOVE "after market" especially if it provides CHEAP solution. I also like your point on customization (which, perhaps, will be more expensive...). From another hand Jonathan's message shows the reality: he bought "after market" aperture (I would think he did specify, which aperture he needs), which does not fit... As I told in my original message, aperture on JEM1200 survived for many years and it was cost to me something like $100 from JEOL. How much I could save on "after market"? $30 perhaps or even less? To me it would be probably easier to get stuff from guaranteed source (JEOL) rather than spent time looking for aperture, which will "fit"... Sergey
At 01:52 PM 3/3/2004, you wrote: } In response to Sergey and Jonathan's questions let me say this: Ladd, as } most people know, produces probably 97% of all apertures made in the U.S. } We customize apertures for all microscopes to fit the user's needs. For } instance in Sergey's case, he could have two or even three 50um holes if he } wanted. The "after market" apertures offer you this customization } opportunity. We can machine in various materials, such as moly, platinum, } etc. and provide strips or discs as thin as 0.025mm. } } Generally, apertures in the "after market" are quite a bit less expensive, } more readily available and, as I mentioned, can be customized to meet your } needs. We put as many as 20 holes in some plates and strips for specialized } equipment. } } As a matter of fact most apertures are "after market" because EM } manufacturers tend not to produce their own apertures. I am not impartial } on this matter, but for flexibility and value the "after market" is the } place to look for apertures. } } Disclaimer: Ladd Research sells EM supplies including apertures and } specialized micro-holes to EM users and microscope manufacturers. } } John Arnott } } Ladd Research } 83 Holly Court } Williston, VT 05495 } } On-line Catalog: http://www.laddresearch.com } } tel: 1-802-658-4961(anywhere) or 1-800-451-3406(US) } fax: 1-802-660-8859 } e-mail: sales-at-laddresearch.com } ----- Original Message ----- } From: "Sergey Ryazantsev" {sryazant-at-ucla.edu} } To: {Microscopy-at-sparc5.microscopy.com} } Sent: Wednesday, March 03, 2004 3:56 PM } Subject: [Microscopy] Re: TEM apertures } } } } } } } } -------------------------------------------------------------------------- } ---- } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -------------------------------------------------------------------------- } ----- } } } } Jonathan } } I do believe, the apertures in JEM1200 are made from molybdenum (or } } tungsten?), definitely not a Pt. Perhaps you have to contact JEOL for } } specification (they are quite friendly). Personally, I would avoid "after } } market" spare parts especially objective apertures (they are designed in } } the way they are actually self-cleaning under the beam). Aperture in my } } microscope is "in" for more than 5 years and I never cleaned it. Deposits } } on the objective aperture usually indicates problem with vacuum. You may } } easily clean original JEOL aperture by heating up in tantalum or tungsten } } boat in good vacuum (5x10-6 torr or better) for a minute or less. It } } cleans everything. The problem there is that it's possible to damage the } } hole itself (well, it means, it's time to buy new one) if overheat. It's } } important, the boat made from different material than aperture, otherwise, } } the strip will stick to the boat. I am using 25-50-100 um objective } } aperture strip. In 95% cases we are using 50um spot - it provides enough } } light with decent contrast. I am using 25um spot (not recommended to use } } with plastic sections!) for very weak W shadowed samples only . I never } } used 100 um. I hope it helps. Sergey } } } } At 11:17 AM 3/3/2004, you wrote: } } } } } } } } --------------------------------------------------------------------------- } --- } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } } To Subscribe/Unsubscribe -- } } } http://www.msa.microscopy.com/MicroscopyListserver } } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } } --------------------------------------------------------------------------- } ---- } } } } } } Greetings: } } } } } } This is a question about TEM objective apertures, mostly about thickness } } } and selection of hole size. } } } } } } We have a JEOL 1200, I wanted to replace the objective aperture, old one } } } was not cleaning up well. Ordered one from EM supplier we get most } } } everything else from. Picked one that was for JEOL, a Pt strip .1 mm } thick, } } } three holes, 50, 100, 150. Sounded perfect. } } } } } } Tried to put it in yesterday. Way too thick. Aperture holder accepts the } } } strip by slipping it in a fixed sandwich kind of space, not with a } separate } } } screw down holder over the top. Old aperture was way thinner than the new } } } one, even though new one was called 'thin'. } } } } } } Checked all the other catalogs I have, most had apertures with the same } } } dimensions. Don't have much experience with this so I am hoping to be } } } educated. } } } } } } Are the apertures for this instrument something special? Where do I get } an } } } aperture to fit? Is JEOL the only source? What criteria do I use to } select } } } the hole size? etc. } } } } } } Help me out on this one. } } } } } } Thanks } } } } } } Jonathan Krupp } } } Microscopy & Imaging Lab } } } University of California } } } Santa Cruz, CA 95064 } } } (831) 459-2477 } } } jmkrupp-at-cats.ucsc.edu } } } } _____________________________________ } } } } Sergey Ryazantsev Ph. D. } } Electron Microscopy } } UCLA School of Medicine } } Department of Biological Chemistry } } 10833 Le Conte Ave, Room 33-089 } } Los Angeles, CA 90095 } } } } Phone: (310) 825-1144 (office) } } (310) 206-1029 (Lab) } } FAX (departmental): (310) 206-5272 } } mailto:sryazant-at-ucla.edu } } } } } } } }
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jvonreis-at-columbiabasin.edu) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, March 3, 2004 at 11:20:34 ---------------------------------------------------------------------------
Email: jvonreis-at-columbiabasin.edu Name: Jennifer von Reis
Question: Does anyone have information on how to use hexamethyldisilazene (HMDS) instead of a critical point drier to prepare specimens, that have been fixed and dehydrated in alcohol, for SEM observation?
We have done quite a bit so far. Most investigations were done on on sidiments ranging between 1m below the surface up to rocks at ~100 m Below the surface. My preference is still to mount in Araldite (the cheap version you can get from fiberglass hobby shops) under vacuum to get rid of trapped gas and polish to 1 micron surface finish. Do not crush. You losse info like pore density and distibution/relationship of the different components. Weathering is more clear in a cross section. Most of the investigation is in BSE mode. Carbon coating is preferred since it interfere less with the EDS spectrum. If you can work in Low Vacuum range (0.1 torr - 1 torr) it helps with reducing charging.
A reasonable refernce is (more suitable to rocks) is "Bachscattred Scanning Electron Microscopy and Image analysis of Sediments and Sedimentary Rocks" by D. H. Kringsly and other authers. Cambridge press ISBN 0-521-45346-1
How can we best prepare our soil samples for SEM and micro analysis? We are novices at this. We have great equipment (Hitachi FESEM 4500 and EDAX Genesis) We are interested in phosphorus and iron in river sediments. We are having difficulty in interpreting the results we have been getting (we crushed soil to double sticky tape, mounted on Al stubs and coated with AuPd). The preps look good at the SEM with little charging.
What we see is grains and some organic matter (OM) and coatings. The spectrum shows some OM and SiO2 along with other particles of mixed composition.
My question is how can we interpret this? What about references etc? We need your expertise!
Your help will be greatly appreciated. Carol Bronick
Agricultural Research Station Box 9061 Virginia State University Petersburg, VA 23806 804.524.6822 cbronick-at-vsu.edu
____________________________________________________________________ Check your SchoolEmail at http://www.CampusI.com Find the LOWEST PRICES on books at http://www.campusi.com/BookFind !
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 00:25:17 2004
-- [ From: Garber, Charles A. * EMC.Ver #3.1a ] --
Jennifer von Reis wrote: ================================================================== Question: Does anyone have information on how to use hexamethyldisilazene (HMDS) instead of a critical point drier to prepare specimens, that have been fixed and dehydrated in alcohol, for SEM observation? ================================================================== A good overview on this topic was published in MICROSCOPY TODAY, May 1997 by Phil Oshel. It has been republished on the SPI Supplies website with permission at URL http://www.2spi.com/catalog/chem/hmds.html
Disclaimer: SPI Supplies is a supplier of HMDS used in electron microscopy applications.
Chuck
============================================
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From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 00:49:29 2004
Sergey; Best way to clean apertures is with a dual beam FIB. That way, you can cut away any dirt, burs, eccentricity, and inspect with the SEM while you are in there!
John Mardinly Intel
-----Original Message----- } From: Sergey Ryazantsev [mailto:sryazant-at-ucla.edu] Sent: Wednesday, March 03, 2004 12:57 PM To: Microscopy-at-sparc5.microscopy.com
Jonathan I do believe, the apertures in JEM1200 are made from molybdenum (or tungsten?), definitely not a Pt. Perhaps you have to contact JEOL for specification (they are quite friendly). Personally, I would avoid "after market" spare parts especially objective apertures (they are designed in
the way they are actually self-cleaning under the beam). Aperture in my
microscope is "in" for more than 5 years and I never cleaned it. Deposits on the objective aperture usually indicates problem with vacuum. You may easily clean original JEOL aperture by heating up in tantalum or tungsten boat in good vacuum (5x10-6 torr or better) for a minute or less. It cleans everything. The problem there is that it's possible to damage the hole itself (well, it means, it's time to buy new one) if overheat. It's important, the boat made from different material than aperture, otherwise, the strip will stick to the boat. I am using 25-50-100 um objective aperture strip. In 95% cases we are using 50um spot - it provides enough
light with decent contrast. I am using 25um spot (not recommended to use with plastic sections!) for very weak W shadowed samples only . I never
used 100 um. I hope it helps. Sergey
At 11:17 AM 3/3/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
A lot of thanks for the information! I am sure that I will contact with you. Thank you -- ************************************************************* Dra. María Belén López Mosquera Unidade de Microscopía Servicios Xerais de Apoio á Investigación Universidade da Coruña Edificio Servicios Centrais de Investigación Campus de Elviña s/n E-15071 A Coruña
O.k., folks I know a number of you are still using TEM film as we are. And due to the pricing of the 250 sheet boxes vs. the 100 sheet boxes you're buying the 250 sheet boxes. Great deal on film cost but the lousy plastic boxes aren't much use.
Those little yellow kodak (or white Ilford) 100 sheet film boxes are great for lots of things but we've been using them for transporting exposed film from the scope rooms to the dark room for processing. I assume that other folks have been doing the same since I've come across this practice at every TEM lab I've ever worked at. But since we've been buying the plastic 250 sheet boxes our supply of the yellow cardboard boxes has been dwindling slowly. SO now the quesiton is what do we replace the 100 sheet cardboard boxes with? Has anyone found the answer yet?
The $40 price premium for buying 100 sheet boxes of film is a bit much for replacing the cardboard boxes withy new cardboard boxes.
The $20K and upwards price for a CCD camera to move to digital is also a bit much.
So before I have my shop folks here make me up some replacment metal boxes, are there any other suggestions out there?
Any of the TEM FIlm vendors out there willing to sell expired 100 sheet film really cheap? (You can even keep the film!)
Thanks.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 08:15:31 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (R.H.Olley-at-reading.ac.uk) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, March 4, 2004 at 03:53:49 ---------------------------------------------------------------------------
Email: R.H.Olley-at-reading.ac.uk Name: Robert H. Olley
Title-Subject: [Microscopy] [Filtered] TEM of Polymers
Question: In answer to the question from Dra. Maria BelÈn LÛpez Mosquera:
One way of looking at polymers, especially hydrocarbon polymers like polypropylene and polyethylene, is to prepare a surface and etch with a permanganic reagent. The etched surface is then either: - replicated and the replica examined under TEM; - gold coated and examined under SEM.
It is very easy to distinguish polyethylene and polypropylene this way.
We have a picture gallery of etched surfaces, albeit for specimens with special thermal or mechanical treatment, on:
----------------------------------- Robert H. Olley Reply to: R.H.Olley-at-reading.ac.uk URL: http://www.personal.rdg.ac.uk/~spsolley -----------------------------------
Boxes that hold 4x5 sheet film work well, although the film slides around a bit. If you know someone who uses a 4x5 camera, you're set. If not, try your local pro film processing lab for empty boxes.
Geoff
Richard Edelmann wrote:
} O.k., folks I know a number of you are still using TEM film as we are. And due to the pricing of the 250 sheet boxes vs. the 100 sheet boxes you're buying the 250 sheet boxes. Great deal on film cost but the lousy plastic boxes aren't much use. } } Those little yellow kodak (or white Ilford) 100 sheet film boxes are great for lots of things but we've been using them for transporting exposed film from the scope rooms to the dark room for processing. I assume that other folks have been doing the same since I've come across this practice at every TEM lab I've ever worked at. But since we've been buying the plastic 250 sheet boxes our supply of the yellow cardboard boxes has been dwindling slowly. SO now the quesiton is what do we replace the 100 sheet cardboard boxes with? Has anyone found the answer yet? } } The $40 price premium for buying 100 sheet boxes of film is a bit much for replacing the cardboard boxes withy new cardboard boxes. } } The $20K and upwards price for a CCD camera to move to digital is also a bit much. } } So before I have my shop folks here make me up some replacment metal } boxes, are there any other suggestions out there? } } Any of the TEM FIlm vendors out there willing to sell expired 100 sheet film really cheap? (You can even keep the film!) } } Thanks. } } } Richard E. Edelmann, Ph.D. } Electron Microscopy Facility Supervisor } 350 Pearson Hall } Miami University, Oxford, OH 45056 } Ph: 513.529.5712 Fax: 513.529.4243 } E-mail: edelmare-at-muohio.edu } http://www.emf.muohio.edu } } "RAM disk is NOT an installation procedure." } } }
-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 08:42:16 2004
I was going to reply, but Chuck already did, sort of. There's also the U. of Florida EM web site "Tips and Tricks": http://www.biotech.ufl.edu/EM/tips/index.html search on How you do the procedure depends a lot on your samples. What are they? Phil
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-- Philip Oshel Supervisor, BBPIC microscopy facility Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 - 1284 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 08:56:07 2004
I have been using the (same) black cardboard box system for some time for transporting film to darkroom! Looks like I will hold onto it. Sorry I don't have any extras; they are in use throughout the lab.
Peggy
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Richard Edelmann [mailto:edelmare-at-MUOHIO.EDU] Sent: Thursday, March 04, 2004 8:23 AM To: microscopy-at-MSA.Microscopy.com
O.k., folks I know a number of you are still using TEM film as we are. And due to the pricing of the 250 sheet boxes vs. the 100 sheet boxes you're buying the 250 sheet boxes. Great deal on film cost but the lousy plastic boxes aren't much use.
Those little yellow kodak (or white Ilford) 100 sheet film boxes are great for lots of things but we've been using them for transporting exposed film from the scope rooms to the dark room for processing. I assume that other folks have been doing the same since I've come across this practice at every TEM lab I've ever worked at. But since we've been buying the plastic 250 sheet boxes our supply of the yellow cardboard boxes has been dwindling slowly. SO now the quesiton is what do we replace the 100 sheet cardboard boxes with? Has anyone found the answer yet?
The $40 price premium for buying 100 sheet boxes of film is a bit much for replacing the cardboard boxes withy new cardboard boxes.
The $20K and upwards price for a CCD camera to move to digital is also a bit much.
So before I have my shop folks here make me up some replacment metal boxes, are there any other suggestions out there?
Any of the TEM FIlm vendors out there willing to sell expired 100 sheet film really cheap? (You can even keep the film!)
Thanks.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 09:22:36 2004
Hello: We have a Reichert-Jung FC4D cryo attachement and are having a problem with some the heaters. We are trying to get through to someone at Leica to help with some questions but in the meantime I was hoping someone might have the name etc of a person we could contact for help. Any help is greatly appreciated because as usual this thing failed just as I have a large backlog of cryo samples. Thanks in advance. Steve
Stephen McCartney Research Associate Materials Research Institute 2108 Hahn Hall Va Tech Blacksburg, VA 24061 540-231-9765 - phone 540-231-8517 - FAX
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 09:31:38 2004
Just check with ANY Photographic/darkroom supply place. Look for something called a Paper Safe. They come in a variety of sizes and range in price from about $13(for 8x10 size that holds about 100 sheets) to around $75(for a DELUXE version). Serve same function as cardboard boxes....just more durable since they are usually made out of ABS plastic See this website for an example(picture) of a Paper Safe ===} http://www.freestylephoto.biz/sc_prod.php?cat_id=1603&pid=1526
Kelly A. Ramos Metallurgical Engineer / Supervisor Argo-Tech Materials Laboratories 23555 Euclid Avenue Cleveland, OH 44117 216-692-5904 or 216-692-5446 (fax) 216-692-5816 http://www.atclabs.com
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 09:42:40 2004
When I bought my TEM 12 years ago, it came with two tin cans. The can look very much like a ordinary can except that the inside of lid has a piece of 10 mm black foam. There is a gap between the side of lid and the foam. It is light tight when the closed.
I think a candy/cooky can of proper size and a piece of black foam inside the lid will do the job.
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I have been using the (same) black cardboard box system for some time for transporting film to darkroom! Looks like I will hold onto it. Sorry I don't have any extras; they are in use throughout the lab.
Peggy
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Richard Edelmann [mailto:edelmare-at-MUOHIO.EDU] Sent: Thursday, March 04, 2004 8:23 AM To: microscopy-at-MSA.Microscopy.com
O.k., folks I know a number of you are still using TEM film as we are. And due to the pricing of the 250 sheet boxes vs. the 100 sheet boxes you're buying the 250 sheet boxes. Great deal on film cost but the lousy plastic boxes aren't much use.
Those little yellow kodak (or white Ilford) 100 sheet film boxes are great for lots of things but we've been using them for transporting exposed film from the scope rooms to the dark room for processing. I assume that other folks have been doing the same since I've come across this practice at every TEM lab I've ever worked at. But since we've been buying the plastic 250 sheet boxes our supply of the yellow cardboard boxes has been dwindling slowly. SO now the quesiton is what do we replace the 100 sheet cardboard boxes with? Has anyone found the answer yet?
The $40 price premium for buying 100 sheet boxes of film is a bit much for replacing the cardboard boxes withy new cardboard boxes.
The $20K and upwards price for a CCD camera to move to digital is also a bit much.
So before I have my shop folks here make me up some replacment metal boxes, are there any other suggestions out there?
Any of the TEM FIlm vendors out there willing to sell expired 100 sheet film really cheap? (You can even keep the film!)
Thanks.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 10:27:48 2004
Dear Richard, My TEM puts the exposed film into a box in the camera, but we found out the hard way that it wasn't quite light tight. I wrap the box in the black plastic bag that the photo paper comes in. These bags work for all types of light-tight transport. Regards, Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 e-mail: mager-at-interchange.ubc.ca ----- Original Message ----- } From: "Richard Edelmann" {edelmare-at-MUOHIO.EDU} To: {microscopy-at-MSA.Microscopy.com} Sent: Thursday, March 04, 2004 5:23 AM
Hi, I need to purchase a number of Philips 6V 10W bayonet style, single contact bulbs for use in the illuminator boxes from Ernst Leitz GMBH Wetzlar transformer housing assemblies. The bulb part number appears to be 6814. Another number that appears on the bulb is k15679. The transformer/bulb housing assemblies are model # 307-127.002. I've searched light bulb web sites and I've tried our local microscope sales rep, no luck on either front. Can anyone out there help?
TIA Mike -- ******************************************************************** Michael M. Cheatham 312 Heroy Geology Laboratory Phone (315)-443-1261 Syracuse University Fax (315)-443-3363 Syracuse, NY 13244-1070
owner of PLASMACHEM-L: http://listserv.syr.edu/archives/plasmachem-l.html owner of XRF-L: http://listserv.syr.edu/archives/xrf-l.html owner of TIMS-L: http://listserv.syr.edu/archives/tims-l.html owner of SIRIS-L: http://listserv.syr.edu/archives/siris-l.html ********************************************************************
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 14:39:06 2004
DISCLAIMER: I have no financial or other interest in the reference quoted above.
Best regards-
David
-- David Henriks Vice President
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Michael Cheatham wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } Hi, } I need to purchase a number of Philips 6V 10W bayonet style, } single contact bulbs for use in the illuminator boxes from Ernst Leitz } GMBH Wetzlar transformer housing assemblies. The bulb part number } appears to be 6814. Another number that appears on the bulb is } k15679. The transformer/bulb housing assemblies are model # 307-127.002. } I've searched light bulb web sites and I've tried our local } microscope sales rep, no luck on either front. Can anyone out there } help? } } TIA } Mike
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 16:40:21 2004
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America To
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 4 17:52:12 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (nagashim-at-ncifcrf.gov) from http://www.msa.microscopy.org/Ask-A-Microscopist.html on Thursday, March 4, 2004 at 11:17:23 ---------------------------------------------------------------------------
Question: I have a problem to mount thin-sections on a thin-bar hexagon grid. The sections do not adhere well and cause many wrinkles on section. There is a glue pen, but I want to know if I can buy solution so I can apply a drop on to grid and let it dry to have a better attachment on.
Richard I do find those plastic boxes quite useful to store the film. As for transportation - I have vacuum desiccator with film set in the dark-room. It's quite convenient: you don't need load-reload films. We are using the rule: each magazine with film should be developed immediately and re-loaded with new film. Nevertheless, for the last two years nobody is willing to use film - we go digital all the time. So, I still keep dark-room in the working order and pay salary to my technician... This is our fate: each new technology adds load to your budget, not relieve. Sergey
At 05:23 AM 3/4/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
My undergrad histology class has given me a new and novel problem. One of the students is experiencing severe motion sickness while she views specimens on a standard binocular compound microscope. I have had students bothered when viewing specimens on the 6-headed scope and someone else is moving the specimen field but this is a first. Any one have an easy suggestion?
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
KUNIO- The best way to get them flat is to make a convex water surface in the boat. That makes the point of contact high where you bring the grid down on the sections and makes the section spread uniformly across the plastic film. It is impossible to prevent all the wrinkles, but this method reduces them to a maneagable number or frequency. Carol
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-- __ Carol A. Heckman, Ph.D. Professor of Biological Sciences Director, Center for Microscopy & Microanalysis Bowling Green State University, Bowling Green, OH 43403 fax: (419) 372-2024 email: heckman-at-bgnet.bgsu.edu ___________________________________________________________________________
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 10:13:08 2004
Hmmm, I remember that I used to get sick while looking at some immunofluorescent slides...I found that if I slowed down (instead of rapidly scanning the slides for areas of interest) and took frequent breaks, I was OK. I would not have called it "severe" motion sickness, though.
When I was on the multi-headed scope with several people and my boss "driving" (i.e. changing magnification, focusing and moving the slide), I learned to anticipate the changes and look away from the scope as the changes were being made.
Not sure if this helps-maybe if she took standard antimotion sickness medicine before getting on the scope would help?
Good luck to you and her- Kathleen Roberts Principal Lab Technician Neurotoxicology Labs Dept of Pharmacology and Toxicology Ernest Mario School of Pharmacy Rutgers University 41 B Gordon Rd Piscataway, NJ 08854
Tom Phillips wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } My undergrad histology class has given me a new and novel problem. One of } the students is experiencing severe motion sickness while she views } specimens on a standard binocular compound microscope. I have had students } bothered when viewing specimens on the 6-headed scope and someone else is } moving the specimen field but this is a first. Any one have an easy } suggestion? } } Thomas E. Phillips, PhD } Professor of Biological Sciences } Director, Molecular Cytology Core } 3 Tucker Hall } University of Missouri } Columbia, MO 65211-7400 } } 573-882-4712 (office) } 573-882-0123 (fax) } PhillipsT-at-missouri.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 10:17:03 2004
I think the easiest, though certainly not the cheapest, solution would be to get either a digital or even a simple video camera mounted on the microscope, then project the live image onto a large monitor or screen. I would bet that with the student's field of view not restricted by looking into eyepieces, the feeling would be greatly reduced. If you coupled this with some type of image capture ability, then you would have the added benefit of being able to provide the students with digital micrographs from their actual session.
Good luck!
David
David Bell Scientist Electron Microscopy Lab Millipore Corporation 80 Ashby Road Bedford, MA 01730 (781) 533-2108
Tom Phillips {phillipst-at-missouri.edu} 03/05/2004 10:16 AM
To: Microscopy-at-msa.microscopy.com cc: Subject: [Microscopy] motion sickness on LM?
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My undergrad histology class has given me a new and novel problem. One of
the students is experiencing severe motion sickness while she views specimens on a standard binocular compound microscope. I have had students
bothered when viewing specimens on the 6-headed scope and someone else is moving the specimen field but this is a first. Any one have an easy suggestion?
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
Tom, My college room mate had that same problem when she was taking Embryology and had to scan slides to find structures. Her way of coping was to take a does of Dramamine about 1/2 hour before her lab class. I had a similar problem, but at the TEM, when I was pregnant. Obviously, a pharmacological solution was out of the question. I found that I could cope with it if I moved the image slowly, glanced away briefly and took a few slow, deep breaths. Your student should make sure that she takes the time to fully adjust the binocular head to her individual needs...set up each eyepiece at focus, so that she can relax her vision. As someone stated last week on this site...she should look THROUGH the microscope, not AT or IN it. That's my 2 cents worth, Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 10:31:56 2004
I'm basing this reply based on explanations and advice from my MD in relation to my own problems with more traditional causes of motion sickness. This is a mixture of the "technical" terms I remember, and the practical explanations that went with them. My doctor is really kind of neat that way.
The basic cause for most motion sickness is a difference in "inputs" from the eyes and the innter ears. In the case of binocular microscopes, both eyes are seeing a moving field, and the ears are saying "we're not moving" and the brain is getting confused. It responds by trying to "cancel out" the stationary input from the ears (vision has priority), and you get dizzy, upset stomache....
This type of case is kind of the inverse of normal motion sickness, where the ears sense movement, but the eyes are not because the surroundings (boat, car interior) are stationary "relative" to the person. (A "special" case of the theory of relativity.)
What the victim needs to do, is take steps to keep the visual stimulus and the inner ear stimulus "in sync". I hate to say it, but she is concentrating too hard on the view in the mircroscope. The user needs to take more frequent and/or longer looks away from the microscope. Looking around the room, and not just at the table may also help. She may also need to get up and walk a little once in a while. Having identified this as a problem for the individual, it is important that she take these steps throughout the session, from the start, not just when she starts feeling poorly. By then it is too late, and brain is starting the process of fighting the confilicting input stimula. It will be an uphill battle to reign things back in.
Another option that may help would be to only use one occular for most viewing, and keep the unused eye open, and looking at something stationary. Those who have used single occular microscopes know that this is not a natural thing to do (at least for the untrained) but it may help balance the stimula.
Different folks react differently to the problem, and need to find a "personalized" solution. This is a game the brain is playing, and it isn't happy about having to do it. Once it starts, it can be difficult to reverse, and an extended break might be the best bet.
I hope this helps.
John W. Raffensperger, Jr.
--- Tom Phillips {phillipst-at-missouri.edu} wrote: } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The } Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } My undergrad histology class has given me a new and } novel problem. One of } the students is experiencing severe motion sickness } while she views } specimens on a standard binocular compound } microscope. I have had students } bothered when viewing specimens on the 6-headed } scope and someone else is } moving the specimen field but this is a first. Any } one have an easy } suggestion? } } Thomas E. Phillips, PhD } Professor of Biological Sciences } Director, Molecular Cytology Core } 3 Tucker Hall } University of Missouri } Columbia, MO 65211-7400 } } 573-882-4712 (office) } 573-882-0123 (fax) } PhillipsT-at-missouri.edu } } }
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 10:33:54 2004
As one who suffers from serious motion sickness I can sympathize with your student.
I think the problem may lie in the oculars not being set up perfectly for her eyes. Another could be that she has sinus problems/inner ear issues and the positioning of her head may be causing imbalance to occur.
Try getting her to fit the oculars better. As we all have experienced when first using LM's not every one know how to optimally set those puppies up. Students are loath to fess up to this and will suffer in silence, or not in the case of your motion sick student.
How this helps,
Paula :-)
Paula Sicurello George Washington Univ. Medical Center Electron Microscope Lab Washington, DC 20037 202-994-2930 phone 202-994-2518 fax } } } Tom Phillips {phillipst-at-missouri.edu} 03/05/04 10:39 AM } } }
------------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
My undergrad histology class has given me a new and novel problem. One of the students is experiencing severe motion sickness while she views specimens on a standard binocular compound microscope. I have had students bothered when viewing specimens on the 6-headed scope and someone else is moving the specimen field but this is a first. Any one have an easy suggestion?
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
Thomas, I do know from experience that this can occur when one is pregnant and when the operator has an ear infection.
Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } My undergrad histology class has given me a new and novel problem. } One of the students is experiencing severe motion sickness while she } views specimens on a standard binocular compound microscope. I have } had students bothered when viewing specimens on the 6-headed scope } and someone else is moving the specimen field but this is a first. } Any one have an easy suggestion? } } Thomas E. Phillips, PhD } Professor of Biological Sciences } Director, Molecular Cytology Core } 3 Tucker Hall } University of Missouri } Columbia, MO 65211-7400 } } 573-882-4712 (office) } 573-882-0123 (fax) } PhillipsT-at-missouri.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 10:48:21 2004
Could there be issues with UV light or ozone from a mercury or xenon lamphouse?
Alan Stone ASTON
At 09:16 AM 3/5/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 14:30:45 2004
} My undergrad histology class has given me a new and novel problem. } One of the students is experiencing severe motion sickness while she } views specimens on a standard binocular compound microscope. I have } had students bothered when viewing specimens on the 6-headed scope and } someone else is moving the specimen field but this is a first. Any } one have an easy suggestion? } On Mar 5, 2004, at 8:44 AM, John Raffensperger wrote:
} The basic cause for most motion sickness is a } difference in "inputs" from the eyes and the innter } ears. In the case of binocular microscopes, both eyes } are seeing a moving field, and the ears are saying } "we're not moving" and the brain is getting confused.
Dear Tom and John, I am not an expert, but I understand that this input mismatch is treated by the brain as a nervous system malfunction like those from ingestion of a toxin, and that the response (motion sickness) has evolved to eliminate the possible toxin--thus the nausea. If the student is having difficulty fusing the two images, this could be another source of input mismatch, so suggest that the student look only with one eye and see if that reduces the problem. Good luck. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 15:26:40 2004
Postdoctoral scientist position Biology Department, Brookhaven National Laboratory Long Island, New York
The position requires a Ph.D. in biophysics, biochemistry, or related field and a strong background in structural biology. Outstanding candidates from physical sciences are also encouraged to apply. The position involves structural study of macromolecular protein complexes by cryo transmission electron microscopy. The ideal candidates should have experience with electron microscopy and computer programming for image process. Facilities include a Jeol 2010FasTEM, a Jeol1200EX, and SGI workstations and a full biochemistry lab.
Interested candidates should send a CV and 3 references to hli-at-bnl.gov
-- Huilin Li Brookhaven National Laboratory Biology Dept. Bldg. 463 Upton, NY 11973 Email: hli-at-bnl.gov phone: (631)344-2931 or (631)344-5066 fax: (631)344-3407 http://www.biology.bnl.gov/structure/li.html
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 15:31:31 2004
The simple solution is to see if she experiences the problem when using only one eye. Monocular scopes work just fine - especially for those of us that have a strongly dominant eye :-)
If you have a videomicroscope setup available, you might try that too.
Have you also considered the possibility that an odor is enhancing the "motion sickness" effect? The few times I have experienced the problem I have been in situations that normally wouldn't bother me - but someone had on a strong perfume/aftershave or had used a vehicle "deodorizer" that was strongly scented. She may be reacting to a combination of scent and sight. If so, either removing the problem odor or having a small fan blow fresh air past her face may help.
- Louise
Louise Harner Research Microscopist Albany International Research Co. 777 West Street, P.O. Box 9114 Mansfield, MA 02048 508-337-9529 phone 508-339-4996 fax Louise_Harner-at-albint.com
= = =
Tom Phillips {phillipst-at-missouri.edu} 03/05/2004 10:16 AM
My undergrad histology class has given me a new and novel problem. One of the students is experiencing severe motion sickness while she views specimens on a standard binocular compound microscope. I have had students bothered when viewing specimens on the 6-headed scope and someone else is moving the specimen field but this is a first. Any one have an easy suggestion?
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
I have advertised Kodak 4489 film on the surplus equipment listing and not have received any inquiries. If you have a Zeiss 9 and need film and/or film racks, please contact me off the listserver. I hate to through this out, but it looks like I must in short order.
Thanks, Ken _______________________________________ Kenneth L. Tiekotter, Adjunct Professor The University of Portland Department of Biology 5000 N Willamette Blvd. Portland, OR 97203 USA
Tel.: 503.943.8861
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 5 19:21:55 2004
NOTICE: THE ZEISS 9s2 USES 7CM X 7CM SIZE FILM, NOT 8.3CM X 10.2CM FILM
I have advertised Kodak 4489 film on the surplus equipment listing and not have received any inquiries. If you have a Zeiss 9 and need film and/or film racks, please contact me off the listserver. I hate to throw this out, but it looks like I must in short order.
Thanks, Ken _______________________________________ Kenneth L. Tiekotter, Adjunct Professor The University of Portland Department of Biology 5000 N Willamette Blvd. Portland, OR 97203 USA
Tel.: 503.943.8861
From MicroscopyL-request-at-ns.microscopy.com Sun Mar 7 10:18:07 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (MBA2038-at-AOL.COM) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, March 7, 2004 at 10:02:25 ---------------------------------------------------------------------------
Email: MBA2038-at-AOL.COM Name: steve yutz
Organization: NONE
Title-Subject: [Microscopy] [Filtered] How much bench work time needs to finish 1kidney, 2 kidney and 3 kidney EM cases without scope from start to finish
Question: i would like to know how much total bench work time needs to finish 1 case of kidny electon microscopy(EM), 2 cases of kidney electon microscopy and 3 cases of kidney microscopy.
Q1. if suppose i wants to do one kidney case of EM, how much total time i have to spent bench work from start to finish without scope.
Q2. if suppose i wants to process two cases of kidney EM together how much total time i have to spent bench work from start to finish without scope.
Q3. if suppose i wants to process three cases of kidne EM together how much total time i have to spent bench work from start to finish without scope.
Q4. how many maximum cases kidney EM can be done per week in 40 hours.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (MBA2038-at-AOL. COM) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Sunday, March 7, 2004 at 10:26:41 ---------------------------------------------------------------------------
Email: MBA2038-at-AOL. COM Name: steve yutz
Organization: NONE
Title-Subject: [Microscopy] [Filtered] Learning EM Programs
Question: i would like to learn Electron microscopy. Is there any program i can go to school in NY, NJ, DE, MD or PA state areas or i can learn on job.
Weill Cornell Medical College is welcoming a new TEM to our facility and the following TEM is available for immediate sale:
JEOL JEM-100CXII 20 - 100kV ASID/TEI Attachment Unit with 4x5 camera accessory 3 Specimen holders 1 Dual specimen holder 1 ASID/TEM specimen holder 1 BSD specimen holder Chiller Extra Filaments 100 Film plates with extra Cassette magazine and receiver All associated support supplies and extra parts that we have on hand.
This microscope has been an excellent performer for us, as we still use it on a daily basis and we are sorry to see it go. It has been under a service contract since it's installation and has had two PM's every year.
Please contact us off list for further specifications and details regarding this great opportunity.
Thanks in advance,
Michael Ganger: mtg2003-at-med.cornell.edu Omayra Velez: omv2001-at-med.cornell.edu Weill Cornell Medical College Department of Pathology Electron Microscopy Laboratory 1300 York Ave New York, NY 10021 212-746-6437
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 8 08:03:27 2004
----- Original Message ----- } From: Carol Heckman {heckman-at-bgnet.bgsu.edu}
Question: Is it possible to strictly confine the illuminated area on a sample to a few square microns (using an aperture) when working in TIRF geometry ?
We use an objektive-based TIRF microscope consisting of an Zeiss Axiovert 200 inverted microscope and an 100x NA1.45 Zeiss objective. To switch to TIRF-mode we tilt a mirror at the entrance of the incident light device (its position is conjugated to the sample plane).
We have a small aperture (rectangular, 2x2microns virtual size on the sample) which is (in epifluorescent wide-field mode) reproduced nicely and sharply on the sample, but when switching to TIRFM angle, the illuminated field seems to blur on one side. Moreover, the illumination of our aperture is no longer homogenous; there seems to be an intensity gradient along the axis from the sharp to the blurry edge.
Sketch:
------------------------- | Our aperture | {- This edge gets blurry. | | | | To reach TIRF geometry, ------------------------- we tilt our laser beam in this direction --} | This Side Seems To get darker.
At www.biophysics.jku.at/bioph/staf/brames/TIRF_problem.htm you can see a comparison between wide-field illumination and TIRF-illumination and you can also see our problem nicely.
So, is there a possibility to image an aperture sharply when using total internal reflection microscopy?
Thanks in advance for your help
Mario Brameshuber
______________________________________ Mario Brameshuber Institute for Biophysics Johannes Kepler University of Linz,Austria Altenbergerstraße 69 4040 Linz
phone: +43/732/2468/9288
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 8 12:05:12 2004
I have a colleague who has an old (1960s vintage?) Zeiss petrographic scope (reflected and transmitted light) which is missing its polarizing lenses. He recently got a price quote on some repairs, including replacement lenses, which comes to several hundreds of dollars -- money he is not ready to spend. To cut costs, the microscope owner is planning to cut polarizers out of a sheet of polaroid film. It seems to me like this shouldn't cause too much of a problem, aside from the possibility that the film might rotate around as he takes the polarizers in and out, but I thought I might put the question out there for people who are more knowledgeable than I. Also, will this cost-cutting really save him much money? How much do polarizing lenses usually cost, anyway?
Thanks,
Peter Selkin pselkin-at-ucsd.edu
Lecturer/Postdoc Scripps Institution of Oceanography, UC San Diego
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 8 15:22:12 2004
I am looking for LM equipment with which to analyze particles on skin, with particular interest in deposition levels. Optimally, I would like to do this with a handheld microscope in the range of 300-1000x. I have a lead or two, but am looking for more options, particularly cost-efficient ones.
If anyone has some leads on companies or ideas for other ways to analyze this microscopically, please contact me. Thanks!
Kirk
Brian (Kirk) Kirkmeyer, Ph.D. Research Scientist, Microscopy International Flavors and Fragrances 1515 State Highway 36 Union Beach, NJ 07735-3542 732-335-2426 / 732-335-2350 FAX brian.kirkmeyer-at-iff.com
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 8 16:07:32 2004
Hi, Generally speaking, polarized light microscopes have polarizing "elements", not lenses. I think there have been a few specialty scopes built in which an actual lens also controls the state of polarization, but the usual tactic is to have flat elements, rather like the film but manufactured to greater tolerance (ie, more uniform surface, better extinction). THe main advantage and I suppose expense is that it is essential to be able to rotate one of these elements. So one of them usually has the mechanical bits to let you rotate (and to read off the angle of rotation). The big problem your colleague will have with the do it your self approach is that without being able to rotate one of the films it will be extremely difficult to get good extinction. On the other hand, if you had the element that could rotate but was missing its optics and you put polarizing film in, that would probably serve quite well.
I would have thought that a few hundred dollars would be fair but many hundreds might be steep. There is always ebay... Tobias
} } } I have a colleague who has an old (1960s vintage?) Zeiss } petrographic scope (reflected and transmitted light) which is } missing its polarizing lenses. He recently got a price quote on some } repairs, including replacement lenses, which comes to several } hundreds of dollars -- money he is not ready to spend. To cut } costs, the microscope owner is planning to cut polarizers out of a } sheet of polaroid film. It seems to me like this shouldn't cause too } much of a problem, aside from the possibility that the film might } rotate around as he takes the polarizers in and out, but I thought I } might put the question out there for people who are more } knowledgeable than I. Also, will this cost-cutting really save him } much money? How much do polarizing lenses usually cost, anyway? } } Thanks, } } Peter Selkin } pselkin-at-ucsd.edu } } Lecturer/Postdoc } Scripps Institution of Oceanography, UC San Diego
We use a sem (cameca Su-30) and have to use epoxy mounts for coarse samples. We can not mount them on aluminium sample holders. We can not put them in sem because of their volume. We have to use epoxy. But is epoxy conductive? I mean a ground problem occurs? Thanks.
Orkun ERSOY Hacettepe University Department of Geological Engineering SEM laboratory
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 04:09:52 2004
What textbooks and multimedia teaching aids for EM in general, SEM and EDS in particular, would you recommend? I am especially looking for those with (at least some) emphasis on low voltage FEG-SEM and low vacuum techinques and theory.
tia, Stefan
+++++++++++++++++++++++++++++++++++++++++++++++++++++++ Dr Stefan Gunnarsson Evolutionsbiologiskt Centrum Evolutionary Biology Centre Enheten för biologisk strukturanalys Microscopy and Imaging Unit Norbyvägen 18A SE75236 Uppsala, Sweden Tel & Fax: +46 - 18 471 2638 +++++++++++++++++++++++++++++++++++++++++++++++++++++++
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 04:53:53 2004
If he buys high quality film it should work well. McCrone Institute sells them for as little as $4 per 2X2 sheet and they have 1/4 an 1/2 wave plates as well.
If you really want to do it rig get come Gnarled lens cement and put glass covers on the plastic filters and mount them in a metal ring to fit the scope. The extra effort will quickly pay off
Gordon Gordon Couger I collect Microscopy links and documentation posted at http://www.couger.com/microscope/links/gclinks.html http://www.science-info.org/ Attributed and anonymous contributions welcome
Tobias Baskin wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } Hi, } Generally speaking, polarized light microscopes have polarizing } "elements", not lenses. I think there have been a few specialty } scopes built in which an actual lens also controls the state of } polarization, but the usual tactic is to have flat elements, rather } like the film but manufactured to greater tolerance (ie, more uniform } surface, better extinction). THe main advantage and I suppose expense } is that it is essential to be able to rotate one of these elements. } So one of them usually has the mechanical bits to let you rotate (and } to read off the angle of rotation). The big problem your colleague } will have with the do it your self approach is that without being able } to rotate one of the films it will be extremely difficult to get good } extinction. On the other hand, if you had the element that could } rotate but was missing its optics and you put polarizing film in, that } would probably serve quite well. } } I would have thought that a few hundred dollars would be fair but } many hundreds might be steep. There is always ebay... } Tobias } } } } } } } } I have a colleague who has an old (1960s vintage?) Zeiss } } petrographic scope (reflected and transmitted light) which is missing } } its polarizing lenses. He recently got a price quote on some repairs, } } including replacement lenses, which comes to several hundreds of } } dollars -- money he is not ready to spend. To cut costs, the } } microscope owner is planning to cut polarizers out of a sheet of } } polaroid film. It seems to me like this shouldn't cause too much of a } } problem, aside from the possibility that the film might rotate around } } as he takes the polarizers in and out, but I thought I might put the } } question out there for people who are more knowledgeable than I. } } Also, will this cost-cutting really save him much money? How much do } } polarizing lenses usually cost, anyway? } } } } Thanks, } } } } Peter Selkin } } pselkin-at-ucsd.edu } } } } Lecturer/Postdoc } } Scripps Institution of Oceanography, UC San Diego } } }
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 05:28:44 2004
The book below is one of the first to have a section on low vacuum/ESEM.
Dave
3rd Edition (2003) Scanning electron microscopy and X-ray microanalysis a text for biologists, materials scientists, and geologists Joseph I. Goldstein ... [et al]
On Tue, 9 Mar 2004 11:28:33 +0100 Stefan Gunnarsson {Stefan.Gunnarsson-at-ebc.uu.se} wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Hi all! } } What textbooks and multimedia teaching aids for EM in general, SEM and } EDS in particular, would you recommend? I am especially looking for } those with (at least some) emphasis on low voltage FEG-SEM and low } vacuum techinques and theory. } } tia, } Stefan } } +++++++++++++++++++++++++++++++++++++++++++++++++++++++ } Dr Stefan Gunnarsson } Evolutionsbiologiskt Centrum Evolutionary Biology Centre } Enheten för biologisk strukturanalys Microscopy and Imaging Unit } Norbyvägen 18A } SE75236 Uppsala, Sweden Tel & Fax: +46 - 18 471 2638 } +++++++++++++++++++++++++++++++++++++++++++++++++++++++ } } } } } This incoming email to UWE has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
This email has been independently scanned for viruses and any virus detected has been removed using McAfee anti-virus software
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 05:30:26 2004
Thanks Richard for his reply and advice. But I doubt about the contact of epoxy and sample holder of SEM (which has 6 holes for aluminium cylinder shaped sample holders). With epoxy there will be no conductance with ground. I will coat the surface of epoxy and sample with carbon but we have to screw it from the body of cylinder shaped epoxy. So do we need to coat the sides of cylinder to make conductance with ground? Or paint the sides of cylinder epoxy with silver?
Orkun ERSOY Hacettepe University Department of Geological Engineering SEM laboratory
-----Original Message----- } From: Richard Beanland [mailto:richard.beanland-at-bookham.com] Sent: 09 Mart 2004 Sali 12:23 To: 'Orkun Ersoy'
Orkun;
There are epoxies that are conductive. For example, there are silver loaded epoxies used in the semiconductor industry to attach microdevices. They are conductive through their bulk and usually have a silver content of } 90% by volume. Epotek and others make these products.
Peter Tomic Agere Systems
-----Original Message----- } From: Orkun Ersoy [mailto:oersoy-at-hacettepe.edu.tr] Sent: Tuesday, March 09, 2004 6:46 AM To: Microscopy-at-MSA.Microscopy.Com
Thanks Richard for his reply and advice. But I doubt about the contact of epoxy and sample holder of SEM (which has 6 holes for aluminium cylinder shaped sample holders). With epoxy there will be no conductance with ground. I will coat the surface of epoxy and sample with carbon but we have to screw it from the body of cylinder shaped epoxy. So do we need to coat the sides of cylinder to make conductance with ground? Or paint the sides of cylinder epoxy with silver?
Orkun ERSOY Hacettepe University Department of Geological Engineering SEM laboratory
-----Original Message----- } From: Richard Beanland [mailto:richard.beanland-at-bookham.com] Sent: 09 Mart 2004 Sali 12:23 To: 'Orkun Ersoy'
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Mike_Steves-at-Pall.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, March 8, 2004 at 14:43:01 ---------------------------------------------------------------------------
Email: Mike_Steves-at-Pall.com Name: Mike Steves
Organization: Pall Corp
Title-Subject: [Microscopy] [Filtered] MListserver: SEM
Question: Is there a stain for cellulose only? we need to see how far it penetrates into a polyethylene substrate, when viewed (X-Sec)in the SEM Would Iodine work? We have the capability to do X-Ray maps as well. Thanks.
For TEM we developed an enzyme-gold label. Details can be found here. Berg, R.H., G.W. Erdos, M. Gritzali and R. Brown 1988. Enzyme-gold affinity labeling of cellulose. ]. EM Tech. 8:371-380
For LM applications, Calcofluor is another option, but requires UV illumination.
At 09:34 AM 3/9/2004, you wrote:
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Gregory W. Erdos Ph.D. Assistant Director, Biotechnology Program Scientific Director, Electron Microscopy P.O. Box 118525 217 Carr Hall University of Florida Gainesville, FL 32611 gwe-at-ufl.edu 352-392-1295 fax- 352-846-0251
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 08:44:16 2004
Hi, For light microscopy, a classic stain is "Calcofluor white" (also known as Cellufluor, and some other names too). This is a fluorescent dye, excited by UV and emitting blue. It will stain other beta 1-4 linked polymers so that in a plant cell wall it is not absolutely specific for cellulose. However, it does stain cellulose brightly and in an artificial composite of cellulose and say polyethylene, it should work perfectly, provided you can accept the lower resolution of the light microscope.
To do this at higher magnification is harder, as far as I know. There are cellulose binding proteins that can be coupled to colloidal gold. These have been used in TEM to detect cellulose, and could probably work in SEM provided you had the magnification to detect the gold particles (typically 5 to 10 nm in diameter). Also I believe that the gold-cellulose binding domain probe is not for sale--you have to make it your self.
Hope this helps, Tobias Baskin
} } Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (Mike_Steves-at-Pall.com) from } http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html } on Monday, March 8, 2004 at 14:43:01 } --------------------------------------------------------------------------- } } Email: Mike_Steves-at-Pall.com } Name: Mike Steves } } Organization: Pall Corp } } Title-Subject: [Microscopy] [Filtered] MListserver: SEM } } Question: Is there a stain for cellulose only? we need to see how } far it penetrates into a polyethylene substrate, when viewed } (X-Sec)in the SEM } Would Iodine work? } We have the capability to do X-Ray maps as well. } Thanks. } } } ---------------------------------------------------------------------------
I would be extremely reluctant to use iodine on samples to be introduced into a SEM due to the aggressivity of iodine vapor in causing corrosion. In any case, iodine forms a complex with starches which would seem not to be the specificity that you are seeking. Iodine is also well known for adding to unsaturated bonds in organic compounds, adding another potential source of confusion.
If you can work with fluorescence microscopy instead of SEM there are some fluorescent dyes in the stilbene class that are often used for tagging cellulose. "Calcofluor" is one of these that is available from histology suppliers. They work in a manner analogous to fabric "whiteners" in detergent that make the laundry fluoresce.
John Twilley Conservation Scientist
by way of MicroscopyListserver wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Mike_Steves-at-Pall.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Monday, March 8, 2004 at 14:43:01 } --------------------------------------------------------------------------- } } Email: Mike_Steves-at-Pall.com } Name: Mike Steves } } Organization: Pall Corp } } Title-Subject: [Microscopy] [Filtered] MListserver: SEM } } Question: Is there a stain for cellulose only? we need to see how far it penetrates into a polyethylene substrate, when viewed (X-Sec)in the SEM } Would Iodine work? } We have the capability to do X-Ray maps as well. } Thanks. } } ---------------------------------------------------------------------------
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 09:37:02 2004
Buehler, and I am sure others, sells a powdered nickel that can be added to their epoxy, rendering the whole mass conductive, I believe. I made a mount for my Philips XL 20 to hold round epoxy samples. It screws into the stage in place of the pin mount holder.
Ron L
-----Original Message----- } From: Orkun Ersoy [mailto:oersoy-at-hacettepe.edu.tr] Sent: Tuesday, March 09, 2004 6:46 AM To: Microscopy-at-MSA.Microscopy.Com
Thanks Richard for his reply and advice. But I doubt about the contact of epoxy and sample holder of SEM (which has 6 holes for aluminium cylinder shaped sample holders). With epoxy there will be no conductance with ground. I will coat the surface of epoxy and sample with carbon but we have to screw it from the body of cylinder shaped epoxy. So do we need to coat the sides of cylinder to make conductance with ground? Or paint the sides of cylinder epoxy with silver?
Orkun ERSOY Hacettepe University Department of Geological Engineering SEM laboratory
-----Original Message----- } From: Richard Beanland [mailto:richard.beanland-at-bookham.com] Sent: 09 Mart 2004 Sali 12:23 To: 'Orkun Ersoy'
Not sure what your specimen stub looks like but I do large metallurgical epoxy embedded specimens all the time. They are mounted in an Aluminum coupon cup and are retained on the SEM stage via standard 3.1mm diameter pin. For a smaller specimen, I would think that the same approach would work.
Sputter coat the embedded specimen. Use a thin, flexible double sticky tab (EMS or Fullam) that just hits the top of the mount and runs down the side. When mounted in the SEM, it will be conductive.
gary g.
At 01:46 AM 3/9/2004, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 10:54:11 2004
Though I don't work with it much these days, I was lucky enough to have taken the "Polarized Light Microscopy" course taught by Mr. McCrone here at work. As far as the lenses go, there aren't polarizing lenses, but there are lenses that are specially made for polarized light microscopy. They are a special variety of achromats called "strain-free". From the McCrone book: "They are carefully made, from the slow cooling of the glass to the final assembly of the lens components, so as to avoid causing strains in the glass which would become visible when used in polarized light microscopy." (*) There are also strain free condenser lenses.
The other suggestions for making the polarizer and analyzer are great, I just thought some may be interested in the lens information.
* Polarized Light Microscopy McCrone, McCrone, & Delly Pub: McCrone Research Institute ISBN 0-250-40262-9
Regards, Darrell
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 17:30:53 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (StAmourOwl-at-charter.net) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 9, 2004 at 13:33:24 ---------------------------------------------------------------------------
Email: StAmourOwl-at-charter.net Name: Rick Dreiling
Question: I am trying to purchase a bottle to store Osmium in. I am running into difficulty about how to clean the bottle. I have found recommendations of cleaning the glass with 5 % Hydrofluroic acid or 30% Nitric Acid. I am not too keen on either of these methods.
Has anyone been able to find a vendor that sells bottles already cleaned?
How do others deal with cleaning the bottles prior to storing Osmium in it? Is their another method I am unaware of?
If you forgo this cleaning step how long does your Osmium last?
I am analysing polished sections containing small (3 -10µm), globular, calcium aluminium oxide inclusions in a steel matrix. Naturally the analysis totals are high as the surrounding steel influences my oxide analysis. Is there a way/formula of determining how much metallic steel is being measured and thus removing it from the results (even though there may be up to 4% iron oxide present)? I have noticed that the amount of iron present is almost equal to the amount that the total exceeds 100. Is it that simple? Does the presence of metallic steel in the analysis affect the ratio of oxides present?
Thank you,
Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 21:25:14 2004
I do metallographic specimens on a somewhat routine basis. For best results, these are epoxy embedded and polished.
If this is done, my EDAX Genesis EDS will do a superior job on analyzing the specimen. The analysis can be a simple ZAF or a more complex PhiZAF or PhiRhoZAF. In either event, the results are very good. The key to obtaining valid quant data is to achieve low Intensity Ratio errors. The EDAX Genesis EDS system helps you do this.
For other systems, I do not know. For stainless steel varieties, I routinely find 0.5-1.5% Fe without problem. Perhaps your collection system is not congruent with your specimens?
gary g.
At 07:26 PM 3/9/2004, you wrote:
} Dear listers, } } I am analysing polished sections containing small (3 -10µm), globular, } calcium aluminium oxide inclusions in a steel matrix. Naturally the } analysis totals are high as the surrounding steel influences my oxide } analysis. Is there a way/formula of determining how much metallic steel } is being measured and thus removing it from the results (even though there } may be up to 4% iron oxide present)? I have noticed that the amount of } iron present is almost equal to the amount that the total exceeds 100. Is } it that simple? Does the presence of metallic steel in the analysis } affect the ratio of oxides present? } } Thank you, } } Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 21:38:01 2004
Possibly you are right about the collection system, we have a kevex detector and some good 'Oxford type' software provided by a local (Australian) genius. What I want to be able to do is discrimnate between the iron from the surrounding steel and the iron present as iron oxide in the non-metallic inclusion, using the resources I have.
-----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] Sent: Wednesday, 10 March 2004 2:41 PM To: Reynolds, Jodi JI Cc: MSA listserver
I have no financial interest in EDAX whatsoever. I am a totally satisfied user. Granted, their software is very complex and quite intense.
I don't claim to be an expert with it. However.... I'm not all that shabby with it. Their HPD feature is very powerful for identifying true and false peaks.
You might send me a specimen and I can run a complete analysis on it. It does not really take all that long.
As with any EDS specimen, it should be polished, flat and non-conductive. I can fix the conductive aspect with coating. So, don't despair in this regard.
Perhaps the EDAX could be a baseline for you relative to your Oxford. Dunno. I have not used that brand nor any others. Thus, the EDAX could be a single data point. But, it really works. IMO.
Again--big disclaimer. No financial interest in EDAX. Just a super satisfied customer.
gary g.
At 07:54 PM 3/9/2004, you wrote: } Possibly you are right about the collection system, we have a kevex } detector and some good 'Oxford type' software provided by a local } (Australian) genius. What I want to be able to do is discrimnate between } the iron from the surrounding steel and the iron present as iron oxide in } the non-metallic inclusion, using the resources I have. } } -----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] } Sent: Wednesday, 10 March 2004 2:41 PM } To: Reynolds, Jodi JI } Cc: MSA listserver } Subject: [Microscopy] Re: SEM metallography } } } I'm not sure that I understand your problem. } } I do metallographic specimens on a somewhat } routine basis. For best results, these are } epoxy embedded and polished. } } If this is done, my EDAX Genesis EDS will do } a superior job on analyzing the specimen. } The analysis can be a simple ZAF or a more } complex PhiZAF or PhiRhoZAF. In either } event, the results are very good. The } key to obtaining valid quant data is to } achieve low Intensity Ratio errors. The } EDAX Genesis EDS system helps you do this. } } For other systems, I do not know. For stainless } steel varieties, I routinely find 0.5-1.5% Fe } without problem. Perhaps your collection } system is not congruent with your specimens? } } gary g. } } } } At 07:26 PM 3/9/2004, you wrote: } } } } Dear listers, } } } } I am analysing polished sections containing small (3 -10µm), globular, } } calcium aluminium oxide inclusions in a steel matrix. Naturally the } } analysis totals are high as the surrounding steel influences my oxide } } analysis. Is there a way/formula of determining how much metallic steel } } is being measured and thus removing it from the results (even though there } } may be up to 4% iron oxide present)? I have noticed that the amount of } } iron present is almost equal to the amount that the total exceeds 100. Is } } it that simple? Does the presence of metallic steel in the analysis } } affect the ratio of oxides present? } } } } Thank you, } } } } Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 23:02:16 2004
I can show you spectra that look totally good and very valid-looking but are totally wrong. This is not known until applying the EDAX HPD feature. The curve does not follow the spectra peaks. This means that the elements in the list are wrong. Plus, the Intensity Ratio values are also likely too huge. Again, a big indicator of wrong data.
A quick, dumb, analysis will result in a seemingly accurate result of element x and y and z. However, one or more of them are not actually there. HPD is the differentiation about this. I deal with Si and Al, W, Ti, Os, Hf, Fe, Ta, et. al. and have to deal with pile up at low eV. EDAX HPD makes sense out of this. The ability to deconvolve is quite powerful.
Ordinarily, an EDS analysis may look OK, but in reality, it is flat wrong. For example a nice peak that shows Fe at 705eV (L alpha) turns out to be F at 677eV (K alpha). So this is the issue of false versus true peaks. It can get very complex and very difficult without software assistance.
It is really identifying real elements versus wrong elements. Easy to do if not careful.
gary g.
At 09:02 PM 3/9/2004, you wrote: } What do you mean by 'true and false' peaks? My samples are polished and } flat but why non-conductive? Shouldn't the surface be able to conduct so } that any surface charge can be drawn away and reduce flaring? Hence } carbon coating? } } Jodi. } } -----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] } Sent: Wednesday, 10 March 2004 3:51 PM } To: Reynolds, Jodi JI } Cc: MSA listserver } Subject: [Microscopy] RE: SEM metallography } } } I have no financial interest in EDAX whatsoever. } I am a totally satisfied user. Granted, their } software is very complex and quite intense. } } I don't claim to be an expert with it. However.... } I'm not all that shabby with it. Their HPD } feature is very powerful for identifying } true and false peaks. } } You might send me a specimen and I can run } a complete analysis on it. It does not really } take all that long. } } As with any EDS specimen, it should be polished, } flat and non-conductive. I can fix the conductive } aspect with coating. So, don't despair in this } regard. } } Perhaps the EDAX could be a baseline for you } relative to your Oxford. Dunno. I have not } used that brand nor any others. Thus, the EDAX } could be a single data point. But, it really works. } IMO. } } Again--big disclaimer. No financial interest } in EDAX. Just a super satisfied customer. } } gary g. } } } At 07:54 PM 3/9/2004, you wrote: } } Possibly you are right about the collection system, we have a kevex } } detector and some good 'Oxford type' software provided by a local } } (Australian) genius. What I want to be able to do is discrimnate between } } the iron from the surrounding steel and the iron present as iron oxide in } } the non-metallic inclusion, using the resources I have. } } } } -----Original Message----- } } From: Gary Gaugler [mailto:gary-at-gaugler.com] } } Sent: Wednesday, 10 March 2004 2:41 PM } } To: Reynolds, Jodi JI } } Cc: MSA listserver } } Subject: [Microscopy] Re: SEM metallography } } } } } } I'm not sure that I understand your problem. } } } } I do metallographic specimens on a somewhat } } routine basis. For best results, these are } } epoxy embedded and polished. } } } } If this is done, my EDAX Genesis EDS will do } } a superior job on analyzing the specimen. } } The analysis can be a simple ZAF or a more } } complex PhiZAF or PhiRhoZAF. In either } } event, the results are very good. The } } key to obtaining valid quant data is to } } achieve low Intensity Ratio errors. The } } EDAX Genesis EDS system helps you do this. } } } } For other systems, I do not know. For stainless } } steel varieties, I routinely find 0.5-1.5% Fe } } without problem. Perhaps your collection } } system is not congruent with your specimens? } } } } gary g. } } } } } } } } At 07:26 PM 3/9/2004, you wrote: } } } } } } } Dear listers, } } } } } } I am analysing polished sections containing small (3 -10µm), globular, } } } calcium aluminium oxide inclusions in a steel matrix. Naturally the } } } analysis totals are high as the surrounding steel influences my oxide } } } analysis. Is there a way/formula of determining how much metallic steel } } } is being measured and thus removing it from the results (even though there } } } may be up to 4% iron oxide present)? I have noticed that the amount of } } } iron present is almost equal to the amount that the total exceeds 100. Is } } } it that simple? Does the presence of metallic steel in the analysis } } } affect the ratio of oxides present? } } } } } } Thank you, } } } } } } Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 23:07:05 2004
Depending on your specimen, it may or may not be conductive. For normal high vacuum work, it needs to be conductive. Now, depending on what the specimen actually is, and ignoring this, I use Au/Pd or Pt coating. At 50A thickness, this shows up as a very tiny peak in the EDS spectra. I just click them out of the analysis for quant.
I do not use Carbon. Too messy. Os is better but VP is ideal.
gary g.
At 09:02 PM 3/9/2004, you wrote: } What do you mean by 'true and false' peaks? My samples are polished and } flat but why non-conductive? Shouldn't the surface be able to conduct so } that any surface charge can be drawn away and reduce flaring? Hence } carbon coating? } } Jodi. } } -----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] } Sent: Wednesday, 10 March 2004 3:51 PM } To: Reynolds, Jodi JI } Cc: MSA listserver } Subject: [Microscopy] RE: SEM metallography } } } I have no financial interest in EDAX whatsoever. } I am a totally satisfied user. Granted, their } software is very complex and quite intense. } } I don't claim to be an expert with it. However.... } I'm not all that shabby with it. Their HPD } feature is very powerful for identifying } true and false peaks. } } You might send me a specimen and I can run } a complete analysis on it. It does not really } take all that long. } } As with any EDS specimen, it should be polished, } flat and non-conductive. I can fix the conductive } aspect with coating. So, don't despair in this } regard. } } Perhaps the EDAX could be a baseline for you } relative to your Oxford. Dunno. I have not } used that brand nor any others. Thus, the EDAX } could be a single data point. But, it really works. } IMO. } } Again--big disclaimer. No financial interest } in EDAX. Just a super satisfied customer. } } gary g. } } } At 07:54 PM 3/9/2004, you wrote: } } Possibly you are right about the collection system, we have a kevex } } detector and some good 'Oxford type' software provided by a local } } (Australian) genius. What I want to be able to do is discrimnate between } } the iron from the surrounding steel and the iron present as iron oxide in } } the non-metallic inclusion, using the resources I have. } } } } -----Original Message----- } } From: Gary Gaugler [mailto:gary-at-gaugler.com] } } Sent: Wednesday, 10 March 2004 2:41 PM } } To: Reynolds, Jodi JI } } Cc: MSA listserver } } Subject: [Microscopy] Re: SEM metallography } } } } } } I'm not sure that I understand your problem. } } } } I do metallographic specimens on a somewhat } } routine basis. For best results, these are } } epoxy embedded and polished. } } } } If this is done, my EDAX Genesis EDS will do } } a superior job on analyzing the specimen. } } The analysis can be a simple ZAF or a more } } complex PhiZAF or PhiRhoZAF. In either } } event, the results are very good. The } } key to obtaining valid quant data is to } } achieve low Intensity Ratio errors. The } } EDAX Genesis EDS system helps you do this. } } } } For other systems, I do not know. For stainless } } steel varieties, I routinely find 0.5-1.5% Fe } } without problem. Perhaps your collection } } system is not congruent with your specimens? } } } } gary g. } } } } } } } } At 07:26 PM 3/9/2004, you wrote: } } } } } } } Dear listers, } } } } } } I am analysing polished sections containing small (3 -10µm), globular, } } } calcium aluminium oxide inclusions in a steel matrix. Naturally the } } } analysis totals are high as the surrounding steel influences my oxide } } } analysis. Is there a way/formula of determining how much metallic steel } } } is being measured and thus removing it from the results (even though there } } } may be up to 4% iron oxide present)? I have noticed that the amount of } } } iron present is almost equal to the amount that the total exceeds 100. Is } } } it that simple? Does the presence of metallic steel in the analysis } } } affect the ratio of oxides present? } } } } } } Thank you, } } } } } } Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 9 23:30:43 2004
Aaahh. Thank you, I can see how that would be a problem with a certain combination of elements. I am lucky in that respect as my sample is not entirely unknown. The inclusions are literally manufactured and the presence of elements which over lap with say Mg, Al and Si the low eV range is highly unlikely, if not totally impossible. Also, since my K alpha iron line (the lement of interset) also overlaps with some highly unlikely candidates (other than manganese and then I'd see the Mn K alpha line too) I am quite confident in my element selection.
Jodi.
-----Original Message----- } From: Gary Gaugler [mailto:gary-at-gaugler.com] Sent: Wednesday, 10 March 2004 4:18 PM To: Reynolds, Jodi JI Cc: MSA listserver
Dear listers,
is there anybody who knows a web-based or other description of EMSA-spectra file format standard?
A full description of the standard, including example formats, can be found at: http://www.amc.anl.gov/ANLSoftwareLibrary/02-EMMPDL/Xeds/EMMFF/Emmff.Total
David Vowles Electron Microscope Unit Dept of Materials Science and Metallurgy University of Cambridge Pembroke St Cambridge UK CB2 3QZ Tel: +44 (0)1223 334325 Fax: +44 (0)1223 334567 Email: djv23-at-cam.ac.uk
On Wed, 10 Mar 2004, Frank Eggert wrote:
} } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Dear listers, } } is there anybody who knows a web-based or other description of } EMSA-spectra file format standard? } } Thanks in advance } } Frank Eggert } } } =========================================== } http://www.microanalyst.net } =========================================== } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 05:31:05 2004
thank you, that was, what I was searching, but not found.
Frank
David Vowles wrote:
} Frank, } } A full description of the standard, including example formats, can be } found at: } http://www.amc.anl.gov/ANLSoftwareLibrary/02-EMMPDL/Xeds/EMMFF/Emmff.Total } } David Vowles } Electron Microscope Unit } Dept of Materials Science and Metallurgy } University of Cambridge } Pembroke St Cambridge } UK CB2 3QZ } Tel: +44 (0)1223 334325 } Fax: +44 (0)1223 334567 } Email: djv23-at-cam.ac.uk } } On Wed, 10 Mar 2004, Frank Eggert wrote: } } }
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 07:10:23 2004
We just store ours in brown glass bottles. All we do in rinse the bottle out with distilled water and wrap parafilm around the lid. Osmium stored this way lasts for weeks without any problem.
Paula :-)
Paula Sicurello George Washington Univ. Medical Center Electron Microscope Lab Washington, DC 20037 202-994-2930 phone 202-994-2518 fax
} } } by way of MicroscopyListserver {StAmourOwl-at-charter.net} 03/09/04 06:46PM } } }
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (StAmourOwl-at-charter.net) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 9, 2004 at 13:33:24 ---------------------------------------------------------------------------
Email: StAmourOwl-at-charter.net Name: Rick Dreiling
Question: I am trying to purchase a bottle to store Osmium in. I am running into difficulty about how to clean the bottle. I have found recommendations of cleaning the glass with 5 % Hydrofluroic acid or 30% Nitric Acid. I am not too keen on either of these methods.
Has anyone been able to find a vendor that sells bottles already cleaned?
How do others deal with cleaning the bottles prior to storing Osmium in it? Is their another method I am unaware of?
If you forgo this cleaning step how long does your Osmium last?
All I do is rinse out the jar with the buffer I'm going to use and then make up my Osmium. I don't bother doing alot of cleaning and haven't had any problems.
I did have a problem when I accidentally used a brown bottle that had been used for osmium to make up a batch of uranyl acetate. I couldn't figure out why the sections would jump off the grid (and run away for dear life!) whenever I put them in my UA solution...live and learn!
Karen Bovard EM Lab Pathology Creighton University Medical Center Omaha, NE
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 07:54:09 2004
Does anyone have any trick/tips that they would share about promoting resin infiltration.
I work in a clinical pathology lab and time is of the essence.
I had read a small blurb in a book about someone putting their straight resin and sections during infiltration under a 100 watt light bulb for the heat to make the resin less viscous. Is this an OK thing to do? I crave real life tips as to what others are doing!
Another question...What do most people feel is the most important step in resin infiltration---the straight resin step vs the step with the resin mixed with propylene oxide? Which step should I be placing the most emphasis on to keep my time in the infiltration step as short as possible and yet get the best possible results. I'm interested in hearing others theories.
(I use EMBed 812 epoxy resin, PO, and process soft tissue for clinical diagnosis--tumors, kidney, muscle, nerve, etc.)
Karen Bovard EM Lab Pathology Creighton University Medical Center Omaha, NE
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 08:05:35 2004
Quite a bit of my work involves looking at inclusions in steel using both metallography and SEM. Since inclusions are small, you're sure to pick up the surrounding steel matrix in the spectra from secondary effects of the electron beam. I usually just ignore it.
If you must determine the iron content within the inclusion, there is an accepted technique I've read about but not personally done. You can deeply etch the surrounding metal matrix with dilute nital (4-10% nitric acid in alcohol) so the inclusions stand proud. Then apply softened replicating (acetate) tape. Let harden, and remove the tape. This should hopefully pull off the inclusions for EDS analysis without the surrounding steel matrix. The inclusions should not react to the nital.
Stu Smalinskas, P.E. Sr. Metallurgist SKF USA Plymouth, Michigan (734) 414-6862
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Jodi wrote:
Dear listers,
I am analysing polished sections containing small (3 -10µm), globular, calcium aluminium oxide inclusions in a steel matrix. Naturally the analysis totals are high as the surrounding steel influences my oxide analysis. Is there a way/formula of determining how much metallic steel is being measured and thus removing it from the results (even though there may be up to 4% iron oxide present)? I have noticed that the amount of iron present is almost equal to the amount that the total exceeds 100. Is it that simple? Does the presence of metallic steel in the analysis affect the ratio of oxides present?
Thank you,
Jodi Reynolds
ReynoldsJ-at-OneSteel.com
__________________________________ Do you Yahoo!? Yahoo! Search - Find what you’re looking for faster http://search.yahoo.com
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 08:07:12 2004
I always used soap and water, followed by several rinses with distilled water. Why does a bottle for osmium need special cleaning? I certainly would not use HF, it will etch the glass.
Geoff
by way of MicroscopyListserver wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 08:16:24 2004
Good morning, Karen, When I worked in a Clinical EM Lab we found that placing our specimens, in 100% resin, into a vacuum oven set at 37 degrees C for an hour helped tremendously. The heat and vacuum together work much better than leaving the vial on the rotator. Also, for large nerve pieces, you could leave them on the rotator overnight in the 1:2 PO:resin then place them under vacuum the next morning. That helped to prevent many of the holes in the axons. As to which step is most important, the PO & resin or 100% resin, they are equally important. The PO removes the alcohol or acetone, as well as making the resin less viscous to pull it into the tissue. But the 100% resin, especially under vacuum, draws out the remaining PO and further infiltrates the tissues. If you have any PO left behind your blocks will not polymerize properly nor cut well. I hope this helps. Good luck! Feel free to write with any other concerns.
Donna R. Clarkson Northrop Grumman Information Technology for U S Army Medical Research Detachment at Brooks City-Base Phone (210) 536-1416 FAX (210) 536-1449 e-mail donna.clarkson-at-brooks.af.mil "Our Army at War--Relevant and Ready"
-----Original Message----- } From: kbovard-at-creighton.edu [mailto:kbovard-at-creighton.edu] Sent: Wednesday, March 10, 2004 8:10 AM To: microscopy-at-ns.microscopy.com
Does anyone have any trick/tips that they would share about promoting resin infiltration.
I work in a clinical pathology lab and time is of the essence.
I had read a small blurb in a book about someone putting their straight resin and sections during infiltration under a 100 watt light bulb for the heat to make the resin less viscous. Is this an OK thing to do? I crave real life tips as to what others are doing!
Another question...What do most people feel is the most important step in resin infiltration---the straight resin step vs the step with the resin mixed with propylene oxide? Which step should I be placing the most emphasis on to keep my time in the infiltration step as short as possible and yet get the best possible results. I'm interested in hearing others theories.
(I use EMBed 812 epoxy resin, PO, and process soft tissue for clinical diagnosis--tumors, kidney, muscle, nerve, etc.)
Karen Bovard EM Lab Pathology Creighton University Medical Center Omaha, NE
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 08:26:20 2004
I think the fastest and easiest way to promote resin infiltration is to use a microwave system, although it does require additional equipment. Microwaves can take you from fresh tissue to polymerized blocks in 4-6 hours, with equal or sometimes better ultrastructure. They are ideal for diagnostic work, because they make the procedure very fast, compared to conventional processing methods, and do not (in my experience) compromise quality. You can find details and protocols for these systems on the websites of vendors selling them, such as Ted Pella and Electron Microscopy Sciences and probably others.
Let me know if you have any questions. I have no financial interest in any of these systems or companies, but our lab has been a very happy user of microwave products.
Good luck.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility---We Do Small Well! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
-----Original Message----- } From: kbovard-at-creighton.edu [mailto:kbovard-at-creighton.edu] Sent: Wednesday, March 10, 2004 8:10 AM To: microscopy-at-ns.microscopy.com
Does anyone have any trick/tips that they would share about promoting resin infiltration.
I work in a clinical pathology lab and time is of the essence.
I had read a small blurb in a book about someone putting their straight resin and sections during infiltration under a 100 watt light bulb for the heat to make the resin less viscous. Is this an OK thing to do? I crave real life tips as to what others are doing!
Another question...What do most people feel is the most important step in resin infiltration---the straight resin step vs the step with the resin mixed with propylene oxide? Which step should I be placing the most emphasis on to keep my time in the infiltration step as short as possible and yet get the best possible results. I'm interested in hearing others theories.
(I use EMBed 812 epoxy resin, PO, and process soft tissue for clinical diagnosis--tumors, kidney, muscle, nerve, etc.)
Karen Bovard EM Lab Pathology Creighton University Medical Center Omaha, NE
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 08:40:18 2004
I don't do a lot of high volume embedding, so I purchase the sealed vials of 4% OsO4 fixative. I then make up small batches of 2% OsO4 (-at-20ml) at a time. I use 20 ml scintillation vials and store it in the metal container that the vials come in.
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: StAmourOwl-at-charter.net [mailto:StAmourOwl-at-charter.net] Sent: Tuesday, March 09, 2004 6:47 PM To: microscopy-at-ns.microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (StAmourOwl-at-charter.net) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 9, 2004 at 13:33:24 ---------------------------------------------------------------------------
Email: StAmourOwl-at-charter.net Name: Rick Dreiling
Question: I am trying to purchase a bottle to store Osmium in. I am running into difficulty about how to clean the bottle. I have found recommendations of cleaning the glass with 5 % Hydrofluroic acid or 30% Nitric Acid. I am not too keen on either of these methods.
Has anyone been able to find a vendor that sells bottles already cleaned?
How do others deal with cleaning the bottles prior to storing Osmium in it? Is their another method I am unaware of?
If you forgo this cleaning step how long does your Osmium last?
Sorry! I failed to mention that once we use up the OsO4, we rinse the scint. vial and discard it in "hazardous" waste container. We don't bother with cleaning and reusing the vials.
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Paula Sicurello [mailto:patpxs-at-gwumc.edu] Sent: Wednesday, March 10, 2004 8:24 AM To: StAmourOwl-at-charter.net; microscopy-at-ns.microscopy.com
We just store ours in brown glass bottles. All we do in rinse the bottle out with distilled water and wrap parafilm around the lid. Osmium stored this way lasts for weeks without any problem.
Paula :-)
Paula Sicurello George Washington Univ. Medical Center Electron Microscope Lab Washington, DC 20037 202-994-2930 phone 202-994-2518 fax
} } } by way of MicroscopyListserver {StAmourOwl-at-charter.net} 03/09/04 06:46PM } } }
---------------------------------------------------------------------------- -- The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (StAmourOwl-at-charter.net) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 9, 2004 at 13:33:24 ---------------------------------------------------------------------------
Email: StAmourOwl-at-charter.net Name: Rick Dreiling
Question: I am trying to purchase a bottle to store Osmium in. I am running into difficulty about how to clean the bottle. I have found recommendations of cleaning the glass with 5 % Hydrofluroic acid or 30% Nitric Acid. I am not too keen on either of these methods.
Has anyone been able to find a vendor that sells bottles already cleaned?
How do others deal with cleaning the bottles prior to storing Osmium in it? Is their another method I am unaware of?
If you forgo this cleaning step how long does your Osmium last?
We use a standard glass bottle with a good plastic screw cap - specifically, it is a Schott brand bottle that one can buy from most major scientific supply houses. These bottles are the ones with blue or orange caps. We start with a new clean bottle and rinse it well with distilled water before using. We reuse the same bottle multiple times (} 1 year usage per bottle) but the caps tend to become black with osmium so we replace those about every 6 months. We keep this bottle in a plastic secondary containment jar. The other jar is covered with aluminum foil to minimize the osmium's exposure to light. We keep the whole assembly in a fume hood at room temperature. Despite careful closing of the inner glass jar, the inside of the plastic container goes black with time. This should be a strong warning to those who keep their osmium in a refrigerator. They are slowly leaking osmium into the refrigerator, the rest of its contents and their lab air.
If the osmium stock goes bad (turns black prematurely), I would discard it and probably the bottle rather than waste time cleaning it. If I had a valuable piece of class contaminated with osmium, I would use hydrogen peroxide (H2O2) to remove the osmium. This works great but it is very reactive and significant quantities of osmium should not be mixed with hydrogen peroxide or you may get a violent exothermic reaction.
} ------------------------------------------------------------------------------- } } Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by } (StAmourOwl-at-charter.net) from } http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, } March 9, 2004 at 13:33:24 } --------------------------------------------------------------------------- } } Email: StAmourOwl-at-charter.net } Name: Rick Dreiling } } Organization: SIUE- Dental School } } Title-Subject: [Microscopy] [Filtered] MListserver: } } Question: I am trying to purchase a bottle to store Osmium in. I am running } into difficulty about how to clean the bottle. I have found } recommendations of } cleaning the glass with 5 % Hydrofluroic acid or 30% Nitric Acid. I am } not too } keen on either of these methods. } } Has anyone been able to find a vendor that sells bottles already cleaned? } } How do others deal with cleaning the bottles prior to storing Osmium in } it? Is } their another method I am unaware of? } } If you forgo this cleaning step how long does your Osmium last? } } ---------------------------------------------------------------------------
Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
We have found that teflon bottles work well with osmium. There are some that are inert to nearly all chemicals, and they don't break if dropped. We wrap the top in parafilm for storage. I just rinse between refills and have no trouble with contamination. Mary Gail Engle
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 09:46:12 2004
I believe that your question is " how many of the x-rays are coming from the steel matrix and not from the inclusion." There are some software applications that address beam interaction area, including Electron Flight Simulator (http://www.small-world.net/efs.htm) (no financial interest, etc, etc.). These would help determine whether the volume of x-ray emission is within a specific size of inclusion. You can also make some estimates from simple nomographs that estimate electron penetration depth as a function of accelerating voltage and sample density. These nomographs can be found in texts and on periodic tables provided by some EDS systems providers.
From my experience, there are other considerations that need to be made on analyzing metallographically polished samples. The first is smearing or carryover of matrix material into the inclusion. Inclusions are often porous, so will capture polishing debris that is removed from the surrounding matrix. Some of the iron that you detect could be polishing residue if you are using mechanical polishing. Polishing compounds can contaminate the inclusions in the same fashion, so you may detect higher silicon or aluminum concentrations if you use silica or alumina polishing compounds. I typically stick to diamond polishing for samples where good EDS data is needed.
Be cautious if you are using a variable pressure SEM. VP is great for analysis of nonconductive samples, but many operators are not aware that the VP environment spreads the beam. The result is that you get significant x-ray production from a much wider area than you would for high vacuum operation. So, analysis of inclusion in steel would show higher iron concentrations with VP than with high vac conditions. Electron flight simulator can produce simulations of beam scatter for various conditions.
Stu's suggestions on separating the inclusions from the matrix is a good one, but you may still have some residue of steel matrix on the inclusions pulled out on the replica.
Hope this helps.
-- Larry D. Hanke, P.E. Materials Evaluation and Engineering, Inc. Practical Solutions Through Technology and Innovation http://www.mee-inc.com (763) 449-8870
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 09:50:40 2004
I store osmium in washed / rinsed scintillation vials with Parafilm wrapped around the vial cap. The vials are stored in a larger jar along with vegetable oil osmium traps. The lid of the outer jar is also sealed with Paraflm. So far I have never had the outermost Parafilm darken since the oil inside the second jar traps any osmium vapors that might get past the Parafilm on the scintillation vial.
Lee Hadden, Ph. D. Department of Biology Wingate University Wingate, NC 28174 704-233-8236
-----Original Message----- } From: Geoff McAuliffe [mailto:mcauliff-at-umdnj.edu] Sent: Wednesday, March 10, 2004 12:23 PM To: by way of MicroscopyListserver Cc: microscopy-at-ns.microscopy.com
I always used soap and water, followed by several rinses with distilled water. Why does a bottle for osmium need special cleaning? I certainly would not use HF, it will etch the glass.
Geoff
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-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
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From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 10:27:01 2004
David Vowles already pointed you to the right place. While you are there, you might want to look at a couple of utilities that I wrote. The descriptions and the links are below:
EELS PLOT, A Windows-based electron energy loss spectroscopy and X-ray energy dispersive spectroscopy plotting and processing program for EMSA formatted spectra that will smooth, differentiate, re-color, re-scale, filter, plot, print, annotate, substract background, integrate intensities, display, overlay, and do limited analysis. http://www.amc.anl.gov/ANLSoftwareLibrary/02-EMMPDL/Eels/EELSPlot/
EM Periodic Table, A Windows-based electron energy loss spectroscopy and X-ray energy dispersive spectroscopy program for displaying energy peak and edge values and for determining possible spectral peak overlap lines. http://www.amc.anl.gov/ANLSoftwareLibrary/02-EMMPDL/Xeds/EMPeriodicTable/
The second of these is included in the first program.
Let me know what you think.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472 Walck-at-PPG.com (412) 820-8651 (office) (412) 820-8515 (fax)
-----Original Message----- } From: Frank Eggert [mailto:Eggert-at-mikroanalytik.de] Sent: Wednesday, March 10, 2004 5:23 AM To: microscopy-at-MSA.Microscopy.com
Dear listers,
is there anybody who knows a web-based or other description of EMSA-spectra file format standard?
You make a good point about the beam spread in a VP SEM. When I'm doing EDS in our ESEM, I normally turn the chamber vapour pressure down to 0 (well, it's not really 0, but it's substantially reduced) during the acquisition. This causes the image to "fade to black", but that's not normally an issue. At least it keeps the beam spread to a minimum.
F.C. Thomas Geological Survey of Canada (Atlantic) Dartmouth, Nova Scotia
-----Original Message----- } From: Larry Hanke [mailto:hanke-at-mee-inc.com] Sent: Wednesday, March 10, 2004 12:02 PM To: MSA listserver Cc: Reynolds, Jodi JI
Jodi:
I believe that your question is " how many of the x-rays are coming from the steel matrix and not from the inclusion." There are some software applications that address beam interaction area, including Electron Flight Simulator (http://www.small-world.net/efs.htm) (no financial interest, etc, etc.). These would help determine whether the volume of x-ray emission is within a specific size of inclusion. You can also make some estimates from simple nomographs that estimate electron penetration depth as a function of accelerating voltage and sample density. These nomographs can be found in texts and on periodic tables provided by some EDS systems providers.
From my experience, there are other considerations that need to be made on analyzing metallographically polished samples. The first is smearing or carryover of matrix material into the inclusion. Inclusions are often porous, so will capture polishing debris that is removed from the surrounding matrix. Some of the iron that you detect could be polishing residue if you are using mechanical polishing. Polishing compounds can contaminate the inclusions in the same fashion, so you may detect higher silicon or aluminum concentrations if you use silica or alumina polishing compounds. I typically stick to diamond polishing for samples where good EDS data is needed.
Be cautious if you are using a variable pressure SEM. VP is great for analysis of nonconductive samples, but many operators are not aware that the VP environment spreads the beam. The result is that you get significant x-ray production from a much wider area than you would for high vacuum operation. So, analysis of inclusion in steel would show higher iron concentrations with VP than with high vac conditions. Electron flight simulator can produce simulations of beam scatter for various conditions.
Stu's suggestions on separating the inclusions from the matrix is a good one, but you may still have some residue of steel matrix on the inclusions pulled out on the replica.
Hope this helps.
-- Larry D. Hanke, P.E. Materials Evaluation and Engineering, Inc. Practical Solutions Through Technology and Innovation http://www.mee-inc.com (763) 449-8870
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 11:19:33 2004
Dear Jodi, I had one student that dissolved the steel completely with bromine/ methanol and filtered out the inclusions for analysis, but I agree with Stu that the acetate replica technique is the one to use to separate the inclusions from the matrix and look at them free of polishing artifacts and beam spread. I have done it and it is quite simple. 1. Etch the sample with the appropriate etch. Fairly deep etch is best. 2. Soften cellulose acetate sheet ( 5 or 6 thou thick), cut into a 1cm by 2cm strip, by placing it on a stack of filter papers moistened with acetone. 3. Put a small pool of acetone on the surface to be replicated and place the acetate sheet on the pool of acetone. I find it helps handling if you bend 5 mm of one end of the acetate up as a handle, before you start. 4. Wait half and hour for all the acetone to dry away and the acetate to go hard and carefully pull the acetate away from the metal. Turn it over and fix to a stub. Either gold or carbon coat or examine in variable pressure, since the acetate is non-conductive. You should have nice inclusions sitting on a replica of the etched metal, with only carbon and oxygen in the acetate to interfere. Other considerations: how high is your x-ray take-off angle? High take-off angle detectors see less of the rim of the inclusion, but it is not something you can change in an existing setup. If you find a big inclusion (10 microns), how low does the Fe content go? You might consider tilting the sample towards the EDS detector to raise the take-off angle. I think that totals above 100% are more a result of imprecision in the ZAF routines than an indication of metallic vs. oxide iron. There is no way to tell them apart in EDS. Good luck. Regards, Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 e-mail: mager-at-interchange.ubc.ca ----- Original Message ----- } From: "Reynolds, Jodi JI" {ReynoldsJ-at-onesteel.com} To: {Microscopy-at-msa.microscopy.com} Sent: Tuesday, March 09, 2004 7:26 PM
Dear listers,
I am analysing polished sections containing small (3 -10µm), globular, calcium aluminium oxide inclusions in a steel matrix. Naturally the analysis totals are high as the surrounding steel influences my oxide analysis. Is there a way/formula of determining how much metallic steel is being measured and thus removing it from the results (even though there may be up to 4% iron oxide present)? I have noticed that the amount of iron present is almost equal to the amount that the total exceeds 100. Is it that simple? Does the presence of metallic steel in the analysis affect the ratio of oxides present?
Thank you,
Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 13:30:27 2004
Karen, To answer a few questions: 1. The heat of the lightbulb should not be a problem because I routinely put my embedding dishes on top of my 60 degree oven overnight to warm and thin the epon before putting the dishes into the oven. I remember when I started in TEM (1971) that our procedure called for putting the molds into 2 ovens, the first was maybe 35 degrees and the second at 60C the next morning.
2.Decades ago a fellow tech. was doing a fast sample prep of melanoma tumors. He needed to section the day after receiving the tissue so he trimmed the samples as small as possible and then used a magnetic stirrer - "flea" real small one, in a scintillation vial to keep the solutions moving all the time. This worked up to the complete epon exchange which was really too dense for the flea to move in. Of course all the times were shortened so that the samples went into the oven by the end of the day. If I remember correctly the oven that he used was set at 70C.
For years I have put difficult samples on a rotating table instead of the table top since I do not have one of those tissue rotators. The motion helps to mix the small amount of solution that remains in the vials into the newly added chemicals.
Can you purchase a microwave oven?
Pat Connelly U of P Biology Dept. Philadelphia PA psconnel-at-sas.upenn.edu * Quoting kbovard-at-creighton.edu: ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } Does anyone have any trick/tips that they would share about promoting } resin infiltration. } } I work in a clinical pathology lab and time is of the essence. } } I had read a small blurb in a book about someone putting their straight } resin and sections during infiltration under a 100 watt light bulb for the } heat to make the resin less viscous. Is this an OK thing to do? I crave } real life tips as to what others are doing! } } Another question...What do most people feel is the most important step in } resin infiltration---the straight resin step vs the step with the resin } mixed with propylene oxide? Which step should I be placing the most } emphasis on to keep my time in the infiltration step as short as possible } and yet get the best possible results. I'm interested in hearing others } theories. } } (I use EMBed 812 epoxy resin, PO, and process soft tissue for clinical } diagnosis--tumors, kidney, muscle, nerve, etc.) } } Karen Bovard } EM Lab } Pathology } Creighton University Medical Center } Omaha, NE
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 14:25:45 2004
SPURR resin is much less dense than Epon. I do believe, you really could short infiltration time with Spurr without compromising the quality. This is total improvisation (never tried!!!!!!!!!!!!!!!!!!!!): for really small pieces (0.5x0.5x0.5mmm) 30-50-70-95% Et-OH 10 min each 2x100% Et-OH - 20 min each Prop Oxide (PO) - 20 min PO:Spurr 1:1 - 40 min on rotator Spurr - 1 h on rotator (tube should be open) fresh Spurr - 1 h on rotator (tube should be open) fresh Spurr in the mold, polymerization at 60oC (not higher), 24 h
Microwave is also good (possible best) idea.
Let me know if it works. Sergey
At 11:46 AM 3/10/2004, you wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
I'm searching for a preparation technology to make a grain size determination with a SEM. The grains should be distributed in a one-layer. Gritting the powder is not satisfying, because the powders (about 1 to 30µm) agglomerates. Does anyone have an idea how to make the preparation? The SEM-picture should be evaluated with an image analysing software.
This is a great idea and I will definitely give it a go.
Cheers,
Jodi.
-----Original Message----- } From: Kestutis Smalinskas [mailto:smalinskas-at-yahoo.com] Sent: Thursday, 11 March 2004 1:22 AM To: microscopy-at-sparc5.microscopy.com; Reynolds, Jodi JI
Hi Larry,
I generally, but not always, use an ultra sonic bath to clean the samples before analysis, do you think this is effective? I'd like to try the acetate tape method and pull the inclusions out, but you're right about the possibility of the incusions taking some of the steel with them. This would be obvious straight away I imagine and perhaps some of the inclusions would be free of steel matrix residue.
Thank you for your help,
Jodi.
-----Original Message----- } From: Larry Hanke [mailto:hanke-at-mee-inc.com] Sent: Thursday, 11 March 2004 3:02 AM To: MSA listserver Cc: Reynolds, Jodi JI
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (maloneyb-at-fiu.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, March 10, 2004 at 09:40:19 ---------------------------------------------------------------------------
Email: maloneyb-at-fiu.edu Name: Barbara
Organization: FCAEM/FIU
Title-Subject: [Microscopy] [Filtered] TEM receipes for T4 bacteriophages
Question: Dear Group: would you please recommend the following: 1. buffers to use especially for T4 bacteriphages 2. what fixatives to use and postfixative 3. what negative stain works best Let me know ASAP thanks for your help Barb
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Mukundan.Chakrapani-at-nrc.ca) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, March 10, 2004 at 13:00:34 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] AFM MAC mode imaging
Question: I have been using Molecular Imaging AFM in the MAC mode to image lipid bilayers under liquid. Everytime I change to a new tip, the images are okay to begin with but the quality deteriorates subsequently. Can anyone comment on this phenomenon?
} ......................could someone explain to me how using a higher } accellerating voltage could/would/should decrease chromatic } aberations in the EM? Particluarly in the TEM.
Richard,
It's as Dave Barnard wrote. The effective spread of focus in the TEM is the chromatic aberration coefficient times the sum of (the relative energy spread in the beam, the relative HT variation during the image acquisition time, and the relative variation in the lens current during the image acquisition time). Increasing the accelerating voltage lowers the relative energy spread term (the deltaE/E term).
The focal length of the objective lens is proportional to the beam energy (higher energy makes the electrons heavier and the lens less able to deflect them) and inversely proportional to the square of the lens current (more current means stronger lens with stronger focusing).
So a small variation in beam energy produces a small variation in the focal length: delF/F is proportional to delE/E. Similarly, a small variation in lens current produces a small variation in the focal length: delF/F is proportional to -2 x delI/I (the 2 x comes from the square and the - comes from the inverse). Where F, E and I are the focal length, beam energy, and lens current, and delF, delE, and delI are their variations.
The constant of proportionality is Cc, the chromatic aberration coefficient. The beam energy variations come from the beam energy spread plus variations in the high-tension power supply. Beam energy spread can be as low as 0.7 eV FWHH for a FEG TEM. HT variation can be 1 part per million root-mean-square. Converting the 0.7 eV full-width half-height to root-mean-square gives 0.7/2.355 = 0.3 eV. Then, for 300kV the relative beam energy spread is 0.3 eV / 0.3 = 1.0 ppm rms. For a 200kV TEM, the relative beam energy spread would be 0.3 eV / 0.2 = 1.5 ppm rms. However, the total 200kV beam variation is not 50% greater than the 300kV result because the HT variation of 1 ppm rms must be added. Since the energy spread and HT variation are independent* we add in quadrature to get sqrt [ 1.0**2 + 1.0**2 ] = 1.4 for 300kV and sqrt [ 1.0**2 + 1.5**2 ] = 1.8 for 200kV. So going to 300kV lowers the beam variation to about 80% of its 200kV value -- this 27% improvement will of course vary depending on the actual beam energy spread and HT variation. .
We then need to add in the contribution coming from the lens current variation. Then we multiply by Cc to get the spread of focus. Again we add in quadrature. Assuming the lens variation is 0.8 ppm rms, and the Cc is 1.5 mm (typical values), we get: Spread of focus = Cc times sqrt [ (delE/E)**2 + (2 x delI/I)**2 ] = 1.5 x sqrt [1.4**2 + 1.6**2] = 3.2 nm at 300kV, and 1.5 x sqrt [1.8**2 + 1.6**2] = 3.6 nm at 200kV. Now the improvement is only 13% on going from 200kV to 300kV. The amount of improvement on increasing from 200 to 300 kV will vary depending on both the relative HT variation and the relative lens current variation.
* "the energy spread and HT variation are independent" -- mostly, see “Estimation of the Electron Beam Energy Spread for TEM Information Limit”, Michael A. O’Keefe, Peter C. Tiemeijer and Maxim V. Sidorov, Microscopy and Microanalysis 8 (2002) supplement 2: 480-481.
That is all. Mike O'Keefe
David Barnard wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } I am thinking that the energy spread of the the source should be the } same reguardless of accellerating voltage and the accellerator adds } the exact same elctron volts to all electrons so the energy } (chromatic) spread arriving at any lens should be the same no matter } the accelerating voltage. Chromatic aberation, however, is a function } of the relative energy spread compared to the final ev. The } deflection error due to the spread at the source has a much smaller } contribution to the total deflection in a higher voltage TEM lens. } } My 2 ev worth } } Dave } } -- } David Barnard } Wadsworth Ctr } NYS Dept Health } Albany NY 12201-0509 } barnard-at-wadsworth.org } 518 473-5299 voice } 518 474-7992 fax
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 10 21:40:05 2004
I'm posting this question on behalf of a colleague:
I would like to buy a PC (Mac or IBM clone) program for drawing stereograms that can show the relative orientation of two phases Eg matrix/precipitate relationships. I have been using Desktop Microscopist but it is an old version and not really compatible with the current operating systems we use.
I would be most grateful of any advice/suggestions you would care to make.
Yours in hope
Kath
Mark Blackford Materials and Engineering Science, ANSTO PMB 1, Menai, N.S.W., 2234 Australia
Phone 61 2 9717 3027 Fax 61 2 9543 7179
Disclaimer: The views expressed in this E-mail message do not necessarily represent the official views of ANSTO from which this message was conveyed.
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 01:20:07 2004
For disaggregation this would be helpful. 50 g aliquots of sample must be treated with about 100 ml dilute HCl for 5 min. in an ultrasonic bath. One liter of distilled water is added to further dilute the acid and left 24 hours to let the particles settle. After drainage of dilute acid, samples are dried overnight at 150 C.(Wohletz et al. 1995- JVGR v. 67)
-----Original Message----- } From: Timo Junker [mailto:timojunker-at-holografie.com] Sent: 10 Mart 2004 Çarsamba 23:35 To: Microscopy-at-MSA.Microscopy.Com
Hi Timo,
I think some indication of the material it is you are trying to disperse would be helpful. Otherwise, any offered solutions may be fruitless.
Regards,
David Bell Scientist Electron Microscopy Lab Millipore Corporation 80 Ashby Road Bedford, MA 01730 (781) 533-2108
"Orkun Ersoy" {oersoy-at-hacettepe.edu.tr} 03/11/2004 02:34 AM
To: {Microscopy-at-MSA.Microscopy.Com} cc: Subject: [Microscopy] RE: powder preparation technology for Grain size determination (SEM) - need help
------------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
For disaggregation this would be helpful. 50 g aliquots of sample must be treated with about 100 ml dilute HCl for 5 min. in an ultrasonic bath. One liter of distilled water is added to further dilute the acid and left 24 hours to let the particles settle. After drainage of dilute acid, samples are dried overnight at 150 C.(Wohletz et al. 1995- JVGR v. 67)
-----Original Message----- } From: Timo Junker [mailto:timojunker-at-holografie.com] Sent: 10 Mart 2004 Çarsamba 23:35 To: Microscopy-at-MSA.Microscopy.Com
I use a technique developed by Millonig:
First, I use Spurr's, after dehydration with acetone, although I am sure that po users culd also use the method. I infiltrate using specimens in microcentrifuge tubes in an old desk-top clinical centrifuge (the one with holders for 4 stnadard 15 ml tubes). Max speed is 2500 rpm (I have no idea what the rcf is).
For each step, I centrifuge for 10 minutes:
5 drops Spurrs in 100 acetone 1:3 Spurr's:acetone 1:1 Spurr's:acetone 3:1 Spurr's acetone 2 changes of 100% acetone Embed in fresh Spurr's.
Using .5 mm cubes of specimen, I can go from water to embedding oven in less than 3 hours.
Best,
Don
______________________________________________________________________ Donald L. Lovett e-mail: lovett-at-tcnj.edu Assoc. Professor, Dept. of Biology voice: (609) 771-2876 P.O. Box 7718 fax: (609) 637-5118 The College of New Jersey Ewing, NJ 08628-0718
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 06:45:56 2004
We are studying the cells of SPUTUM with fluorescence (syto 25). we are continuously having troble with the high autofluorescence of epithelial cells (from mouth). We are using optilyse to treat the cells before staining.
Does anybody know the cause for the autofluorescence and/or the treathent to lower the autofluorescence.
Thanks
Ari Kuusisto Research physicist
PerkinElmer Life and Analytical Sciences tel. +358-2-2678 508 fax. +358-2-2578 357 E-mail: Ari.Kuusisto-at-PerkinElmer.com
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 09:55:23 2004
A while back, someone on the list requested a spare holder for 3.5 x 4 in. film and I believe that it was for a Hitachi scope. I recently found some in a box that had two labels on it, I thought that they were for a Philips but they came from a Hitachi. If they are still needed, I have a bunch of them but would like a good description of the part you wish in case they are not a match.
Please contact me directly.
Pat Connelly The University of Pennsylvania Department of Biology Philadelphia, PA 19104-6018 215-898-7145 psconnel-at-sas.upenn.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 10:09:38 2004
Our facility is in the process of acquiring an Arcturus LCM system. We have been given a preview of both systems by an Arcturus representative. But we are in need of some advice from investigators who have used either the PixCell IIe and/or the AutoPix systems. Any information regarding the advantages or disadvantages of each system would be appreciated. Particular issues that concern us and would appreciate any response on are:
Ease of use Initial set up time for each of the systems Resolution of each system Sampling time for Proteomics and purity of samples for downstream processing for ex. 50 to 500 samples Can the template for the AutoPix be saved for future use? Reliability of both hardware and software Maintenance costs for each system.
Thank you in advance for your feedback.
aruna
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 11:37:57 2004
I'm involved in a project working with gold and silver coins recovered from a shipwreck. The ship went down in the 1800's. I am seeking advice on how to clean some of the coins in a way that will not remove any of the gold or silver from the coins. Some of the coins have coral on them, some have iron oxide, some cupric cloride, and some have iron sulfide on them. The sulfide is the real problem. Do any of you have suggestions as to how to clean the coins chemically? I don't have the equipment to clean them by electrolysis, and cannot polish them, for that would decrease their value. Thank you in advance for your suggestions.
Edward Haller Diagnostic Electron Microscopy Lab University of South Florida Pathology Department Tampa, FL 33612 (813)974-9584
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 11:41:05 2004
The Electron Microscopy Facility-Medical School at the University of Wisconsin is hosting a three-day workshop on immunogold techniques from May 24-26, 2004. Dr. Jan Luenissen from Aurion Immunogold Reagents & Accessories, an internationally known expert in the field, will be the instructor for the workshop. The workshop will include lectures, hands-on training, round table discussions, and presentations on applications. Also, participants of the workshop will be able to work on their own samples during the workshop. The workshop main curriculum is detailed below. If you are interested in attending or need more information about the workshop, please contact the workshop technical coordinator Hong Yi by phone (404-727-8692) or email (hyi-at-emory.edu).
MAIN CURRICULUM
The properties of gold particles and their protein conjugates. Theories underlying immunogold labeling protocols. Silver enhancement of gold particles Immunogold labeling on a variety of sample preparations for LM. Immunogold labeling for EM a. Pre-embedding immunogold labeling using ultrasmall gold conjugates and silver enhancement. b. Post-embedding immunogold labeling using conventional colloidal gold conjugates and ultrasmalll gold conjugates. Pre- and post-embedding double immunogold labeling. Background minimization in immunogold labeling Signal amplification in immunogold labeling.
Thanks and we hope to see you in Madison.
Randall Massey Operations Manager UW-Electron Microscope Facility 1300 University Ave. Madison, WI 53706 voice (608)262-2993 Fax (608)262-7306
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 13:19:30 2004
Hi Karen We have a Pelco Microwave with a coolspot and a vaccum chamber. If we have all the chemicals made up and ready to go (including the resin) we can go from fixing and cutting in 2 hours. The results are as if we had done it conventionally. The resin is an Epon-Spurr mix and usually cuts easier than Epon alone. Elaine
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-- Dr. Elaine Humphrey Director, BioImaging Facility President, Microscopy Society of Canada University of British Columbia 6270 University Blvd, mail-stop Botany Vancouver, BC CANADA, V6T 1Z4 Phone: 604-822-3354 FAX: 604-822-6089 e-mail: ech-at-interchange.ubc.ca website: www.emlab.ubc.ca
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 13:26:52 2004
A grad student working with plant tissue in our labs wants to purchase a diamond knife. Supplier catalogs indicate different specification when you purchase a knife for TEM ultra-structure. When cutting plant tissue I use a "knife angle" 4 degrees an obtain great results. My question concerns the manufactures description of a 35 or 45 degree knife. Is this the angle of the diamond mounted in the boat ? Knowing we plan to cut plant tissues imbedded in spurr, what angle knife would you suggest we select and why (35 or 45) ? If anyone can also give me a good reference for diamond knives and cutting that would be great.
If you wish you can forward your emails directly to dufresne-at-ms.umanitoba.ca
Thanks, André
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 13:39:52 2004
I have a customer that needs to section drilling mud core samples using a cryostat. He is using a tungsten carbide knife and the sections are 30 microns thick. The problem is that no matter what we try, tape or plastic tubing, the section falls apart. He needs to do some quantitative analysis on the specimen.
We have been considering trying to embed it in a plastic resin and section it on a microtome instead. I am not sure which plastic resin would be best in this application. Also can we use plastic with 3 inch diameter cores or would we have to divide the cores into quadrants.
I am hoping some of the materials people may be able to help me get on the right path. Thanks for your help.
Cheryl Rehfeld Meyer Instruments, Inc. 281-579-0342 e-mail csr-at-meyerinst.com
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 14:24:20 2004
I agree with most of the posts presented so far. I did want to offer a few more thoughts on quantitative analysis.
First, it seems like precious few people know how to do good quantitative EDS these days. Or maybe I should say that few people are aware of the many possible pitfalls. Most issues are rather common sense if one stops to think about it; however, it is so darn easy to push the quantify button on an EDS system without thinking about where the numbers come from and what might have happened along the way.
Let me also say up front that I think most EDS manufacturers probably have decent quantitative analysis routines. I don't think the problem is necessarily in the software. It is usually a problem of not using it correctly. Again, systems have become dreadfully easy to use and a novice user can crank out results. Some of them might even be right.
Current PCs offer plenty of CPU storage and memory that allow for all kinds of neat tricks. They can afford the new operator the benefit of a lot of previous expertise. But I don't think that should absolve the new user of the task of learning many of those principles themselves, such as properly identifying the elements present.
You mentioned using "Oxford-type software". I have no idea what that means. I am still using Oxford's ISIS suite of programs, but that doesn't mean much to the EDAX or Noran users out there. You will have to describe what you mean by that.
It seems one main question for your sample is where are the x-rays being generated? You say you might have some Fe in your inclusion which complicates the issue. I agree with the suggestion to use a program like Electron Flight Simulator to model the interaction volume. You might benefit from cutting back on your beam voltage. (You didn't tell us what you were using.) Even if your beam only penetrates 1 um and your inclusion is 3 um in diameter, you don't really know how thick a layer of inclusions you have left on any given inclusion.
You also did not say if you were using spot mode on the center of the inclusion or using a raster. Spot mode might be better in this case as the raster will approach the edge of the inclusion and fluoresce more neighboring Fe.
Your inclusion is definitely non-conductive. If left as is, the inclusion will probably charge and deflect the beam onto the steel around it. Variable pressure SEM was suggested as one possible remedy for dealing with the charging problem and someone already stated that will lead to scattering of the beam and fluorescence of the neighboring Fe. That is probably not a good idea in your case unless you can collect data at a variety of pressures and try to extrapolate back to 0 pressure. The suggestion was also made to coat the sample with something conductive and then ignore that element in the analysis. I have done that, but still you must be careful. I use a layer of evaporated (not sputtered or flashed) C on my standards. I have seen a measurable change in intensity as a result of the C layer. If you are using your analysis total to judge the results, you need to be aware that the coating will change the total.
You may wish to try the following exercise for sake of learning more about your system. Try performing a linescan across a large inclusion. Our ISIS software allows for a quick intensity linescan. A couple of minutes is enough to produce a fairly good scan under our conditions. I make sure to include a background window for reference. In this case, I would set one close to Fe. You could also do point analyses across the interface, but that would take longer. Then look to see how quickly and how much the Fe level drops as you scan across the inclusion. If you reach a flat bottom on the Fe scan then you might be sure that you have avoided influence from the surrounding steel, if not, then you know your x-rays are not from the inclusion only.
You said your totals do not equal 100%. That definitely requires more information. I think most casual users simply select the normalize function so their totals always come out to 100%. You have not done that, but what does your system do for standards and for matrix corrections? If your spectra were not collected under the same conditions (voltage, beam current, coating, geometry, etc) as your standards, then the totals cannot be expected to match. Then there is an issue of having a standard close in composition to your unknown. Errors in the matrix correction can then partially cancel out. However, I do not know the particulars of your system.
I will add that I was caught by surprise trying to analyze a sample which had small Si inclusions in Al. I tried bending the rules to take an overall analysis of the mixture at low magnification. The results were grossly in error. It turned out that there is a strong linkage between Al and Si in the matrix correction, but my sample had little interaction between the two elements in practice. I basically had domains of pure Si and pure Al, yet the matrix correction assumed a homogeneous mixture and calculated accordingly, but in error. I had enabled normalization, but my total would have been far off from 100% even if I was operating at the same beam current. That could have tipped me off that something was wrong, in this case, a lack of homogeneity.
I apologize for the long note, but I hope it is helpful.
Warren
At 09:26 PM 3/9/2004, you wrote:
} Dear listers, } } I am analysing polished sections containing small (3 -10µm), globular, } calcium aluminium oxide inclusions in a steel matrix. Naturally the } analysis totals are high as the surrounding steel influences my oxide } analysis. Is there a way/formula of determining how much metallic steel } is being measured and thus removing it from the results (even though there } may be up to 4% iron oxide present)? I have noticed that the amount of } iron present is almost equal to the amount that the total exceeds 100. Is } it that simple? Does the presence of metallic steel in the analysis } affect the ratio of oxides present? } } Thank you, } } Jodi Reynolds.
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 14:39:21 2004
You can find useful information on manufacturers sites, for example http://www.microstartech.com/ http://www.emsdiasum.com/Diatome/diamond_knives/default.htm
45 degrees knives are used for routine work.
Vladimir M. Dusevich, Ph.D. Electron Microscope Lab Manager 3127 School of Dentistry 650 E. 25th Street Kansas City, MO 64108-2784
} -----Original Message----- } From: Andre Dufresne [mailto:dufresne-at-Ms.UManitoba.CA] } Sent: Thursday, March 11, 2004 1:43 PM } To: microscopy-at-ns.microscopy.com } Subject: [Microscopy] Diamond Knife } } } } } -------------------------------------------------------------- } ---------------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ----------------- } } } A grad student working with plant tissue in our labs wants to } purchase } a diamond knife. Supplier catalogs indicate different specification } when you purchase a knife for TEM ultra-structure. When } cutting plant } tissue I use a "knife angle" 4 degrees an obtain great results. My } question concerns the manufactures description of a 35 or 45 degree } knife. Is this the angle of the diamond mounted in the boat ? } Knowing we plan to cut plant tissues imbedded in spurr, what angle } knife would you suggest we select and why (35 or 45) ? If anyone can } also give me a good reference for diamond knives and cutting } that would } be great. } } If you wish you can forward your emails directly to } dufresne-at-ms.umanitoba.ca } } Thanks, } André } } }
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 15:06:56 2004
I would talk to at least two experts in the field before you try anything. By expert I mean someone with real credentials who works in the field, not some advice you get over the internet. My understanding is that coins are not supposed to be cleaned, period.
Geoff
Edward Haller wrote:
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From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 15:21:29 2004
Your 4 degrees is the clearance angle...the angle at which the entire knife sits in the stage (sets the "attack" angle of the knife edge and that keeps the block from dragging down the back of the knife as it passes). I was taught to set glass knives at 4 degrees, and most of the diamond I've had over the years were designed to work best at 6. The manufacturer's 35 or 45 degrees refers to angle of the bevel of the knife edge itself. I use an ultra-45 for animal-based biological work. I don't know which would be better for plant material. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 11 15:43:56 2004
You are well advised to consult a maritime archaeological conservator before you do anything to the coins.
The last thing archaeologists want is for them to be made bright and shiny.
cheers
rtch
Date sent: Thu, 11 Mar 2004 12:45:22 -0500 } From: Edward Haller {ehaller-at-hsc.usf.edu} Send reply to: ehaller-at-hsc.usf.edu To: microscopy-at-msa.microscopy.com
I have been seeing some of the comments on diamond knives and thought I would share my recent experience with the new vibrating diamond knife from Diatome (i am happy customer with no financial interest). This knife is fantastic. It significantly reduces compression and i get a lot less chatter in some of my hard to cut blocks (either plant seed material or lymphoid tissue). I cut a mix of either Lowicryl K4M, LR Gold and epoxy resin blocks. It works well with all of them. It is a little early to say but I think I am getting less tearing and holes in sections that have poorly infiltrated regions (e.g., starch grains in maize endosperm). The concept of a diamond knife was first proposed by Daniel Studer and he has a paper on it that was published about 3 years ago. So if you have the bucks, consider one of these.
At 04:32 PM 3/11/2004 -0500, you wrote:
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Thomas E. Phillips, PhD Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
André Clearance angle of 4 deg indicates, your particular sample is quite hard to cut. As smaller clearance angle or actual knife angle it's less compression in your sections (better). From this point of view, 35 deg knife should serve you better. From another hand, 35 deg diamond knife is quite fragile: its easier to damage such knife, than "normal" 45 deg knife. If the person, who is going to use knife is novice, I would suggest 45 deg. 35 deg knife is for "profi" in my opinion. I hope it may help. Sergey
At 11:43 AM 3/11/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
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André Clearance angle of 4 deg indicates, your particular sample is quite hard to cut. As smaller clearance angle or actual knife angle it's less compression in your sections (better). From this point of view, 35 deg knife should serve you better. From another hand, 35 deg diamond knife is quite fragile: its easier to damage such knife, than "normal" 45 deg knife. If the person, who is going to use knife is novice, I would suggest 45 deg. 35 deg knife is for "profi" in my opinion. I hope it may help. Sergey
At 11:43 AM 3/11/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
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A grad student working with plant tissue in our labs wants to purchase a diamond knife. Supplier catalogs indicate different specification when you purchase a knife for TEM ultra-structure. When cutting plant tissue I use a "knife angle" 4 degrees an obtain great results. My question concerns the manufactures description of a 35 or 45 degree knife. Is this the angle of the diamond mounted in the boat ? Knowing we plan to cut plant tissues imbedded in spurr, what angle knife would you suggest we select and why (35 or 45) ? If anyone can also give me a good reference for diamond knives and cutting that would be great.
If you wish you can forward your emails directly to dufresne-at-ms.umanitoba.ca
Thanks, André
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 00:35:24 2004
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Hello,
I have a customer that needs to section drilling mud core samples using a cryostat. He is using a tungsten carbide knife and the sections are 30 microns thick. The problem is that no matter what we try, tape or plastic tubing, the section falls apart. He needs to do some quantitative analysis on the specimen.
We have been considering trying to embed it in a plastic resin and section it on a microtome instead. I am not sure which plastic resin would be best in this application. Also can we use plastic with 3 inch diameter cores or would we have to divide the cores into quadrants.
I am hoping some of the materials people may be able to help me get on the right path. Thanks for your help.
Cheryl Rehfeld Meyer Instruments, Inc. 281-579-0342 e-mail csr-at-meyerinst.com
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 03:24:31 2004
Have you looked at any other LASER microdissection systems? We use a PALM set-up and are very pleased with it. See
www.palm-microlaser.com
Note - I have no commercial connection with PALM or any of its associates.
} Our facility is in the process of acquiring an Arcturus LCM system. We } have been given a preview of both systems by an Arcturus } representative. But we are in need of some advice from investigators } who have used either the PixCell IIe and/or the AutoPix systems. Any } information regarding the advantages or disadvantages of each system } would be appreciated. Particular issues that concern us and would } appreciate any response on are: } } Ease of use } Initial set up time for each of the systems } Resolution of each system } Sampling time for Proteomics and purity of samples for downstream } processing for ex. 50 to 500 samples } Can the template for the AutoPix be saved for future use? } Reliability of both hardware and software } Maintenance costs for each system. } } Thank you in advance for your feedback. } } aruna
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 07:09:02 2004
We seem to be a bit hostile to this request. Before we get too upset, we should remember there is a difference between cleaning coins and restoring coins. I have recently rediscovered my childhood hobby of coin collecting and from what I read cleaning coins is still (after more than 30 years of not collecting) a hot topic.
Maybe the best advice this group can give is to provide referrals to experts.
Frank Karl Degussa Corporation Akron Technical Center 3500 Embassy Parkway Suite 100 Akron, Ohio 44333
330-668-2235 Ext. 238
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 07:16:46 2004
Hi all, We had a superconductor named YBCO (Y1Ba2Cu3O7-x)(ceramic)with transition temperature 91 K. We put it on sem without coating and got good result below x17 000 magnifications. I coated it with carbon and now we get good results on x25 000 magnifications (I think the limit of our Sem-Cameca Su-30).
That was a superconductor but we had to coat it. The contamination on the surface of material (which can not be seen with naked eye)may result that. I just wanted to share my experience with you. All interpretations are wellcome.
Orkun ERSOY Hacettepe University Department of Geological Engineering SEM Laboratory
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 14:25:33 2004
My two cents. I have worked with plant materials for many years. Using standard 45 D-knife and UltraCut E Microtome$B!J(BReichert-Jung) I haven$B!G(Bt had any problem with the standard 45 to cut Epon/standard, Epon/Araldit mix, Spur and LR White plastics. I haven$B!G(Bt tried 35 Degree D-Knife and have no right to say which one is better than the other.
Haixin Xu Biological Sciences Univ. of Maryland Baltimore County
white -----Original Message----- } From: Leona Cohen-Gould [mailto:lcgould-at-med.cornell.edu] Sent: 2004$BG/(B3$B7n(B11$BF|(B 16:33 To: Andre Dufresne; microscopy-at-ns.microscopy.com
Your 4 degrees is the clearance angle...the angle at which the entire knife sits in the stage (sets the "attack" angle of the knife edge and that keeps the block from dragging down the back of the knife as it passes). I was taught to set glass knives at 4 degrees, and most of the diamond I've had over the years were designed to work best at 6. The manufacturer's 35 or 45 degrees refers to angle of the bevel of the knife edge itself. I use an ultra-45 for animal-based biological work. I don't know which would be better for plant material. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 15:14:04 2004
} I'm involved in a project working with gold and silver coins recovered } from a shipwreck. The ship went down in the 1800's. I am seeking advice } on how to clean some of the coins in a way that will not remove any of } the gold or silver from the coins. Some of the coins have coral on } them, } some have iron oxide, some cupric cloride, and some have iron sulfide } on } them. The sulfide is the real problem. Do any of you have suggestions } as } to how to clean the coins chemically? I don't have the equipment to } clean them by electrolysis, and cannot polish them, for that would } decrease their value. Thank you in advance for your suggestions. } Dear Edward, You can wash the coins with a mild detergent without harming them, and acetone or xylene can be used to get off grease. Do not use abrasives. Gold is inert to many chemical treatments, but use them only if the milder treatments do not work. Silver is more reactive than gold, so chemical treatment would only be used as a last resort--it would usually be right to leave a foreign material on the coins rather than using chemical treatment to remove it. That said, some corrosion can be dissolved with ammonia. Removing the sulfide is definitely not recommended, since that would definitely decrease the value of the coins. If you can reproduce the features of the contamination on some silver coins of nominal value, you can test the effect of a proposed treatment before using it on a much more valuable specimen. Good luck. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 15:49:58 2004
I have never used any diamond knives for semi-thin plastic sections: 0.5-0.75 microns thick, but I noticed them on the DIATOME web site today, and I wondered what you folks might think of these knives. I really love the idea of getting rid of glass knives once and for all, except perhaps to rough in the blocks.
What do you people think of "histo" diamond knives for semi-thin sectioning?
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 17:27:21 2004
My three cents (the Canadian dollar being chronically lower than the US Greenback):
1. the 35 degree knife is an excellent knife IF USED WITH CARE. This can be as simple as reducing the size of the block face as much as possible.
2. For soft materials, compression is indeed reduced.
3. Perhaps surprisingly, the 35 is much superior to higher knife angles at sectioning 'hard' materials successfully, meaning useable pieces of appropriate thickness. ('Shattering' of the section is common, but the thickness is usually reasonably close to the set thickness, unless ultrathin ( {30 nm sections) are desired).
4. In the materials lab where I spent the greater part of my career, and where we did a lot of contract work, our two 35's were in constant use. The 45s and a 55 collected dust.
Tom
Dr. Tom Malis Scientist Advisor Natural Resources Canada Govt. of Canada Phone: 613-995-7358 cell: 613-371-4577 FAX: 613-947-6606 malis-at-nrcan.gc.ca
-----Original Message----- } From: Haixin Xu To: 'Leona Cohen-Gould'; 'Andre Dufresne'; microscopy-at-ns.microscopy.com Sent: 3/12/2004 10:16 AM
Hi,
My two cents. I have worked with plant materials for many years. Using standard 45 D-knife and UltraCut E Microtome(Reichert-Jung) I haven?t had any problem with the standard 45 to cut Epon/standard, Epon/Araldit mix, Spur and LR White plastics. I haven?t tried 35 Degree D-Knife and have no right to say which one is better than the other.
Haixin Xu Biological Sciences Univ. of Maryland Baltimore County
white -----Original Message----- } From: Leona Cohen-Gould [mailto:lcgould-at-med.cornell.edu] Sent: 2004?3?11? 16:33 To: Andre Dufresne; microscopy-at-ns.microscopy.com
Your 4 degrees is the clearance angle...the angle at which the entire knife sits in the stage (sets the "attack" angle of the knife edge and that keeps the block from dragging down the back of the knife as it passes). I was taught to set glass knives at 4 degrees, and most of the diamond I've had over the years were designed to work best at 6. The manufacturer's 35 or 45 degrees refers to angle of the bevel of the knife edge itself. I use an ultra-45 for animal-based biological work. I don't know which would be better for plant material. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 12 18:10:00 2004
It's just simply great. I used to make huge 1.5 um semithin sections from the whole mouse brain (coronal) and it was nightmare with glass. Then I was trying tungsten carbide - sections were milky and did not stick well to the glass slide (and yes, very scratchy). I think, histo-knife is great for semithin. Sergey
At 02:05 PM 3/12/2004, you wrote:
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Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
I have used "histology" diamond knives very successfully for many years for semi-thin sectioning and for sectioning of hard (materials science) specimens. That being said, such knives are manufactured by several companies, and there can be substantial differences in quality between "brands."
best regards, Steven Slap
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From MicroscopyL-request-at-ns.microscopy.com Sat Mar 13 11:53:19 2004
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Hi Karen We have a Pelco Microwave with a coolspot and a vaccum chamber. If we have all the chemicals made up and ready to go (including the resin) we can go from fixing and cutting in 2 hours. The results are as if we had done it conventionally. The resin is an Epon-Spurr mix and usually cuts easier than Epon alone. Elaine
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-- Dr. Elaine Humphrey Director, BioImaging Facility President, Microscopy Society of Canada University of British Columbia 6270 University Blvd, mail-stop Botany Vancouver, BC CANADA, V6T 1Z4 Phone: 604-822-3354 FAX: 604-822-6089 e-mail: ech-at-interchange.ubc.ca website: www.emlab.ubc.ca
From MicroscopyL-request-at-ns.microscopy.com Sun Mar 14 12:44:35 2004
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Hi
Have you looked at any other LASER microdissection systems? We use a PALM set-up and are very pleased with it. See
www.palm-microlaser.com
Note - I have no commercial connection with PALM or any of its associates.
} Our facility is in the process of acquiring an Arcturus LCM system. We } have been given a preview of both systems by an Arcturus } representative. But we are in need of some advice from investigators } who have used either the PixCell IIe and/or the AutoPix systems. Any } information regarding the advantages or disadvantages of each system } would be appreciated. Particular issues that concern us and would } appreciate any response on are: } } Ease of use } Initial set up time for each of the systems } Resolution of each system } Sampling time for Proteomics and purity of samples for downstream } processing for ex. 50 to 500 samples } Can the template for the AutoPix be saved for future use? } Reliability of both hardware and software } Maintenance costs for each system. } } Thank you in advance for your feedback. } } aruna
From MicroscopyL-request-at-ns.microscopy.com Sun Mar 14 14:19:08 2004
We've used histoknives from both Diatome and Drukker with great success cutting sections from 300 nm up to 2 microns thick and block faces several mm wide. One reservation we had when we first started was the life span of the edge. However this has not been a problem and we find we can get a few years use out of a knife (mainly resin embedded plant material)
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I have never used any diamond knives for semi-thin plastic sections: 0.5-0.75 microns thick, but I noticed them on the DIATOME web site today, and I wondered what you folks might think of these knives. I really love the idea of getting rid of glass knives once and for all, except perhaps to rough in the blocks.
What do you people think of "histo" diamond knives for semi-thin sectioning?
Ian Hallett HortResearch Mt Albert Research Centre, Private Bag 92 169 Auckland, New Zealand Fax +64 9 815 4201 Telephone +64 9 815 4200 EMail ihallett-at-hortresearch.co.nz
______________________________________________________ The contents of this e-mail are privileged and/or confidential to the named recipient and are not to be used by any other person and/or organisation. If you have received this e-mail in error, please notify the sender and delete all material pertaining to this e-mail. ______________________________________________________
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 02:32:40 2004
I got a wide 'histo' diamond when starting a project involving lots of mouse bone work. It is obviously much more forgiving to those in which the decalcification was not quite complete. I roughly trim them all on glass then pop the diamond in to take a good face off the whole batch. I think histo diamonds are a real boon and well worth the money.
Gill Brown Histopathology Group GlaxoSmithKline Medicines Research Centre, Gunnelswood Road, STEVENAGE, Hertfordshire. SG1 2NY tel. +44 (0)1438 764119 fax. +44 (0)1438 764782 email. gillian.2.brown-at-gsk.com
----- Forwarded by Gillian 2 Brown/PharmRD/GSK on 15-Mar-2004 08:37 -----
"Garry Burgess" {GBurgess-at-exchange.hsc.mb.ca}
12-Mar-2004 22:05
To: Microscopy
cc: Subject: [Microscopy] Histo Diamond Knife for Semi-thin sections
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I have never used any diamond knives for semi-thin plastic sections: 0.5-0.75 microns thick, but I noticed them on the DIATOME web site today, and I wondered what you folks might think of these knives. I really love the idea of getting rid of glass knives once and for all, except perhaps to rough in the blocks.
What do you people think of "histo" diamond knives for semi-thin sectioning?
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 07:23:37 2004
I have used vinegar to dissolve calcium from sea cucumber and egg shell. It can be used for cleaning coral on gold coin. For other coins, I don't know. Good luck
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On Mar 11, 2004, at 9:45 AM, Edward Haller wrote:
} I'm involved in a project working with gold and silver coins recovered } from a shipwreck. The ship went down in the 1800's. I am seeking advice } on how to clean some of the coins in a way that will not remove any of } the gold or silver from the coins. Some of the coins have coral on } them, } some have iron oxide, some cupric cloride, and some have iron sulfide
} on } them. The sulfide is the real problem. Do any of you have suggestions
} as } to how to clean the coins chemically? I don't have the equipment to } clean them by electrolysis, and cannot polish them, for that would } decrease their value. Thank you in advance for your suggestions. } Dear Edward, You can wash the coins with a mild detergent without harming them, and acetone or xylene can be used to get off grease. Do not use abrasives.
Gold is inert to many chemical treatments, but use them only if the milder treatments do not work. Silver is more reactive than gold, so chemical treatment would only be used as a last resort--it would usually be right to leave a foreign material on the coins rather than using chemical treatment to remove it. That said, some corrosion can be dissolved with ammonia. Removing the sulfide is definitely not recommended, since that would definitely decrease the value of the coins. If you can reproduce the features of the contamination on some
silver coins of nominal value, you can test the effect of a proposed treatment before using it on a much more valuable specimen. Good luck. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 08:01:55 2004
Garry, I haven't cut anything with glass in YEARS! (except when I'm teaching someone how to section). I wouldn't be without a Histo knife. Period. Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 10:13:19 2004
This is reckless, as copper that is part of the alloys will be attacked and in the presence of salt (these are salt water archeological finds) may form chloride-acetate double salts that later unduce further corrosion or efflorescence problems. The long term results of various cleaning methods have been studied by individuals with experience in handling archaeological finds and the benefits and detriments are known. The cleaning of marine archaeological finds should be left to a professional conservator with experience in dealing with salt contaminated corrosion.
John Twilley Conservation Scientist
Ann-Fook Yang wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } I have used vinegar to dissolve calcium from sea cucumber and egg shell. } It can be used for cleaning coral on gold coin. For other coins, I don't } know. Good luck } } Ann Fook } } } On Mar 11, 2004, at 9:45 AM, Edward Haller wrote: } } } I'm involved in a project working with gold and silver coins } recovered } } from a shipwreck. The ship went down in the 1800's. I am seeking } advice } } on how to clean some of the coins in a way that will not remove any } of } } the gold or silver from the coins. Some of the coins have coral on } } them, } } some have iron oxide, some cupric cloride, and some have iron sulfide } } } on } } them. The sulfide is the real problem. Do any of you have suggestions } } } as } } to how to clean the coins chemically? I don't have the equipment to } } clean them by electrolysis, and cannot polish them, for that would } } decrease their value. Thank you in advance for your suggestions. } }
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 11:29:28 2004
I have been using DDk diamond knives to do cryo-ultra-thin cutting of plastics for AFM phase imaging. I found knife damages developed as soon as cut the first sample. I used 35-D first and then 45-D angle knives and saw the same problem. I think I might need to try a Diatome knife now. Can anybody comment on the qualities between DDK and Diatome knives?
} From: Tom Phillips {phillipst-at-missouri.edu} } To: Microscopy-at-msa.microscopy.com } Subject: [Microscopy] Re: Re: Diamond Knife } Date: Thu, 11 Mar 2004 17:42:37 -0600 } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 11:33:22 2004
John brings to mind a good point. We can't assume that these coins are pure silver or pure gold. Given 1800's metallurgy, I'd be amazed if the coins were "pure anything". The coins could easily contain microinclusions and secondary intermetallic phases which may react badly to any chemical cleaning methods.
Stu Smalinskas Metallurgist
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Edward Haller wrote: I'm involved in a project working with gold and silver coins recovered from a shipwreck. The ship went down in the 1800's. I am seeking advice on how to clean some of the coins in a way that will not remove any of the gold or silver from the coins. Some of the coins have coral on them, some have iron oxide, some cupric chloride, and some have iron sulfide on them. The sulfide is the real problem. Do any of you have suggestions as to how to clean the coins chemically? I don't have the equipment to clean them by electrolysis, and cannot polish them, for that would decrease their value. Thank you in advance for your suggestions.
Ann wrote: I have used vinegar to dissolve calcium from sea cucumber and egg shell. It can be used for cleaning coral on gold coin. For other coins, I don't know. Good luck
Ann Fook
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
John wrote: This is reckless, as copper that is part of the alloys will be attacked and in the presence of salt (these are salt water archeological finds) may form chloride-acetate double salts that later unduce further orrosion or efflorescence problems. The long term results of various cleaning methods have been studied by individuals with experience in handling archaeological finds and the benefits and detriments are known. The cleaning of marine archaeological finds should be left to a professional conservator with experience in dealing with salt contaminated corrosion.
John Twilley Conservation Scientist
__________________________________ Do you Yahoo!? Yahoo! Mail - More reliable, more storage, less spam http://mail.yahoo.com
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 13:31:01 2004
When you take a picture of anatase particles distributed on a carbon film, cells adhering to a substrate, or even crystals on Mars, the resulting image is just the first step toward better understanding of the world. It doesn't matter whether your "picture" is on photographic film, within a digital file, or even presented as some other record of light, electrons, x-rays, or other phenomena, it can and should be analyzed to extract quantitative scientific facts. The picture is the art; the analysis is the science.
The New England Society for Microscopy and the Connecticut Microscopy Society announce that pre-registration has opened for the Image Processing Workshop to be held Thursday April 29th. 2004, at Woods Hole, MA.
Details may be found at {http://www.msa.microscopy.com/MSALAS/NESM/NESMHome.htm} http://www.msa.microscopy.com/MSALAS/NESM/NESMHome.htm
Navigate to "Future Meetings".
Initial details about the NESM/CMS Spring Symposium, to be held following the Image Processing Workshop on April 30th. and May 1st., also at Woods Hole, are also available on the same web site. Full details of the Spring Symposium will be available before the end of March.
********************************************
Anthony J. Garratt-Reed, M.A., D.Phil. Manager, Shared Experimental Facilities Center for Materials Science and Engineering Room #13-1027 Massachusetts Institute of Technology, 77 Massachusetts Avenue Cambridge, Massachusetts 02139-4307 USA
Steve is quite right. Some years ago, we evaluated a number of histos from different companies on a variety of materials (Al alloy, Al-SiC composite, etc) and found that some cut nearly as good as a conventional diamond knife, while others performed poorly. Even the best of them left a lot of fine knife marks, though, sometimes covering the entire section, leading us to wonder if the knife edge was somehow serrated.
On the other hand, there were some pleasant surprises. A year or so later, we had a request to section largish (10-20 micron) particles of an amorphous Fe-Nd-B intermetallic used for magnets. It chewed up the edge of all the conventional knives - 35. 45 and 55 - yet a histo (Diatome, if memory serves) produced decent 30 nm thick sections with no edge damage! Ever since, as Steve notes, we first use a histo on any 'hard' material with which we are unfamiliar.
Finally, in another series of tests, we were able to cut 1 micron semithin sections of Al alloy with no knife damage. Intriguingly, ultrathin sectioning of the same alloy showed no 'curling' of the sections (due to residual stress buildup) that can be such a headache when sectioning metals. No idea why, unless the above hypothetical serrated edge somehow evened out the strain buildup.
So, if one has a variety of materials to section, including many that are 'hard' in nature, investing in a histo might be wise so long as you are prepared for the variability Steve mentions.
Tom
Dr. Tom Malis Scientist Advisor Natural Resources Canada Govt. of Canada Phone: 613-995-7358 cell: 613-371-4577 FAX: 613-947-6606 malis-at-nrcan.gc.ca
-----Original Message----- } From: Steven E. Slap To: Garry Burgess Cc: Microscopy-at-sparc5.microscopy.com Sent: 3/13/2004 11:56 AM
Dear fellow microscopists
I have used "histology" diamond knives very successfully for many years for semi-thin sectioning and for sectioning of hard (materials science) specimens. That being said, such knives are manufactured by several companies, and there can be substantial differences in quality between "brands."
best regards, Steven Slap
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From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 14:17:14 2004
I am looking for a hand tool that could press dry fine powder samples into a tablet with a surface that would be suitable for EDS analysis in an SEM. I have looked at things like KBr pellet makers but not sure that would work for this application. Any ideas would be appreciated.
Roy Beavers
Southern Methodist University Department of Geological Sciences P.O. Box 750395 Dallas, Tx 75275 Voice: 214-768-2756 Fax: 214-768-2701 Email: rbeavers-at-mail.smu.edu
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 16:13:12 2004
There are my 3 canadian cents to this thread (equivalent to 2 US cents).
There is a great FREE software for simulating the interactions "primary electrons -matter" and subsequent XR emission. The program can be downloaded from http://www.gel.usherb.ca/casino/ . This software can simulate multilayered sample as well as bi-materials with an interface perpendicular to the scanned surface. It might be vey useful for studying inclusions in steel.
-----Message d'origine----- De : Reynolds, Jodi JI [mailto:ReynoldsJ-at-OneSteel.com] Envoyé : 10 mars, 2004 16:59 À : Larry Hanke Cc : Microscopy (E-mail) Objet : RE: RE: SEM metallography
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Hi Larry,
I generally, but not always, use an ultra sonic bath to clean the samples before analysis, do you think this is effective? I'd like to try the acetate tape method and pull the inclusions out, but you're right about the possibility of the incusions taking some of the steel with them. This would be obvious straight away I imagine and perhaps some of the inclusions would be free of steel matrix residue.
Thank you for your help,
Jodi.
-----Original Message----- } From: Larry Hanke [mailto:hanke-at-mee-inc.com] Sent: Thursday, 11 March 2004 3:02 AM To: MSA listserver Cc: Reynolds, Jodi JI
Yes, but the point is that by cleaning them you destroy all of the information about what happens to coins in that marine environment.
An experienced marine archaelologist or archaeological conservator should be brought into the project.
cheers
rtch
Date sent: Mon, 15 Mar 2004 09:52:16 -0800 (PST) } From: Kestutis Smalinskas {smalinskas-at-yahoo.com}
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dvanoekelen-at-omnilabo.be) from on Monday, March 15, 2004 at 01:30:08 ---------------------------------------------------------------------------
Email: dvanoekelen-at-omnilabo.be Name: Van Oekelen
Organization: Omnilabo
Title-Subject: [Microscopy] [Filtered] SEM from
Question: Dear Microscopists,
I know there are a lot of specialists regarding the use of SEM. Therefore, I was hoping that someone could help me finding a beautiful SEM image of a salt crystal.
What kind of salt? I have marine salt, table salt and Kosher salt. All look different.
What resolution and what useage?
gary g.
At 03:06 PM 3/15/2004, you wrote:
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From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 17:53:26 2004
We recently removed the cover to do some maintenance on our MT2B microtomes. During the process I noticed that the set screw holding the knob to the pivot control screw was loose. When we reassembled the microtome and began cutting sections it became obvious that the upper thickness control was not in proper alignment. Previously we set the upper thickness at 10 to get silver sections. Not anymore. Does anyone know the procedure for realigning the upper thickness control knob? Or, does any one have a service manual that describes the procedure? You may email me directly or post on the list server.
Bob Robert J. Schmitz Department of Biology University of Wisconsin Stevens Point Stevens Point, WI 54481 715-346-2420 FAX 715-346-3624 Email: rschmitz-at-uwsp.edu http://biology.uwsp.edu/faculty/RSchmitz/Default.htm
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 18:37:27 2004
To all experts out there: I was checking to see if anyone out there has been successful in changing a freon insulated HV tank (ours is a 200 KV JEOL) to SF6 gas (with or without outside assistance). If so, please let me about the success of your experience and what internal/external components you may changed and how it went...and if possible, please offer me any advice in the process. Thanks, Michael Coveillo
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 15 19:35:58 2004
Michael, The National Center for Electron Microscopy {http://ncem.lbl.gov/} converted a JEOL ARM-1000 from freon to SF6 several years ago. Someone there should be able to answer your question. Mike O'Keefe
coviello wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } To all experts out there: } I was checking to see if anyone out there has been successful in } changing a freon insulated HV tank (ours is a 200 KV JEOL) to SF6 gas } (with or without outside assistance). If so, please let me about the } success of your experience and what internal/external components you may } changed and how it went...and if possible, please offer me any advice in } the process. } Thanks, } Michael Coveillo
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 07:44:31 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (kssim-at-mmu.edu.my) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 16, 2004 at 07:33:48 ---------------------------------------------------------------------------
Currently, I am trying to interface JEOL SEM 840A model with DT3152. I appreciate those who have experience to interface SEM with DT3152, on how to tap the signal scan x, sacn y and videos signal and connect to DT3152. Please kindly send email to me and supply me with information.
Someone asked me about how to assess the purity of a preparation of yeast mitochondria on a sucrose gradient. As I have no experience of this myself, I would be very glad if you could help me out. Should we do it with EM or would an approach with fluorescent markers for mitochondria do a better job?
Thanks, Stefan
+++++++++++++++++++++++++++++++++++++++++++++++++++++++ Dr Stefan Gunnarsson Evolutionsbiologiskt Centrum Evolutionary Biology Centre Enheten för biologisk strukturanalys Microscopy and Imaging Unit Norbyvägen 18A SE75236 Uppsala, Sweden Tel & Fax: +46 - 18 471 2638 +++++++++++++++++++++++++++++++++++++++++++++++++++++++
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 09:07:19 2004
List members who have an interest in materials science are cordially invited to a free two-day "mini-workshop" on cryoultramicrotomy for the materials sciences. This topic is of special interest for those who work with polymers or other materials which could benefit from ultrathin sectioning or surface "polishing" at low temperatures (no embedding required).
This invitation is extended to persons who are located in the greater Philadelphia/Baltimore/Washington, D.C./New York City area. The workshop will be hosted by the Center for Advanced Scientific Imaging (CASI) at West Chester University of Pennsylvania.
The details are as follows:
**What**: Mini-Workshop on Cryoultramicrotomy for Materials Science
**When**: Tuesday, March 30, 2004, 11:00 am through Wednesday, March 31, 2004, 4:00 pm.
Visitors are invited to arrive early on Tuesday, March 30 for tours of the Center for Advanced Scientific Imaging. The lab will be open at 8:00 am on Tuesday.
**Where**: West Chester University of Pennsylvania, Schmucker Science Center South, Room SSS017. Schmucker Science Center South is located on the corner of South Church Street and Rosedale Avenue in West Chester, PA. West Chester is located approximately 30 miles from Philadelphia.
**Format**: A presentation on cryoultramicrotomy and its applications in the materials sciences will be given on the first day, as well as demonstrations on the preparation of cryotools (hair probes, large and small wire loops, etc.) and glass knife making and evaluation. The care and cleaning of diamond knives will also be discussed.
The demonstrations will be followed by open lab sessions for the attendees to prepare their own cryotools and glass knives.
Also on the first day an introduction to the cryoultramicrotome will be given, and attendees will have an opportunity for hands-on use of the instrument.
The second day will be reserved for attendees to sign up in small groups for additional time and training on the instrumentation, depending on the attendees' individual needs.
**Limited Space Available**: The lectures and demonstrations on the first day are open to everyone, but space is limited to *10 persons* for the in-depth training and extended use of the instrumentation on the second day.
**Contacts**: To RSVP or to reserve a seat for the second day's sessions, please contact any of the following people:
Dr. Fred Monson, CASI / West Chester University of Pennsylvania, 610.738.0437, {fmonson-at-wcupa.edu}
Dr. Robert Chiovetti, RMC Products / Boeckeler Instruments, Inc., 520.745.0001, {bob-at-boeckeler.com}
I have recently encountered a problem I've not dealt with before. I'm putting cover slips on thin sections of geode materials using Permount. When first covered, they are fine, but within 24 hours they develop tiny grains in the Permount, but only over certain mineral grains (usually quartz grains that have inclusions of anhydrite). I'm stumped as to what may be causing this. My only recourse at present is to clean off the cover slip, re-mount and photograph while the Permount is fresh. Any suggestions as to how to deal with this so it does not occur will be welcome.
Henry Barwood Associate Professor of Science, Earth Science Department of Math and Physics MSCX 312G Troy State University Troy, Alabama 36082 hbarwood-at-troyst.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 12:00:31 2004
This may be a stupid question, but there's no way that that an electron beam of say, 15 or 20KeV can hurt a diamond in any way is there? I was checking out a gemstone (unofficially - off the clock :-)for a colleague, which turned out to be a cubic zirconium, but then I got thinking - if it had been a diamond (carbon in crystal form)would that have been beam-sensitive in the way that organic (carbon-based) things are? My hunch is no, but if somebody ever asks me to check out their engagement ring to see if it's really diamond (yes, I hear people sometimes do that ;-), I don't want to be the guy that burns a hole in it.
Frank
F.C. Thomas FThomas-at-NRCan.gc.ca, 902-426-4635, facsimile 902-426-6152 Natural Resources Canada, Bedford Institute of Oceanography, P.O. Box 1006, Dartmouth, Nova Scotia B2Y 4A2 Ressources naturelles Canada, l'Institut Oceanographique du Bedford, B.P. 1006, Dartmouth, (Nouvelle-Ecosse) B2Y 4A2 Government of Canada/Gouvernement du Canada
-----Original Message----- } From: dvanoekelen-at-omnilabo.be [mailto:dvanoekelen-at-omnilabo.be] Sent: Monday, March 15, 2004 7:07 PM To: microscopy-at-ns.microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dvanoekelen-at-omnilabo.be) from on Monday, March 15, 2004 at 01:30:08 ---------------------------------------------------------------------------
Email: dvanoekelen-at-omnilabo.be Name: Van Oekelen
Organization: Omnilabo
Title-Subject: [Microscopy] [Filtered] SEM from
Question: Dear Microscopists,
I know there are a lot of specialists regarding the use of SEM. Therefore, I was hoping that someone could help me finding a beautiful SEM image of a salt crystal.
EDX will unlikely tell you anything about the difference between a synthetic diamond and a naturally occurring one, just in case you were thinking about going into the jewelry business. Zr is another matter, since it's elementally different than diamond, man made or natural. Regarding your question, I cannot imagine a beam current high enough in an SEM that would alter a diamond, especially with it's high thermal conductivity. Some new man-made gems are virtually indistinguishable from natural gems. It makes the diamond mining people very nervous.
I'm sure you'll hear from others on this subject.
Peter Tomic Agere Systems
-----Original Message----- } From: Thomas, Frank [mailto:FThomas-at-NRCan.gc.ca] Sent: Tuesday, March 16, 2004 1:19 PM To: microscopy-at-ns.microscopy.com
This may be a stupid question, but there's no way that that an electron beam of say, 15 or 20KeV can hurt a diamond in any way is there? I was checking out a gemstone (unofficially - off the clock :-)for a colleague, which turned out to be a cubic zirconium, but then I got thinking - if it had been a diamond (carbon in crystal form)would that have been beam-sensitive in the way that organic (carbon-based) things are? My hunch is no, but if somebody ever asks me to check out their engagement ring to see if it's really diamond (yes, I hear people sometimes do that ;-), I don't want to be the guy that burns a hole in it.
Frank
F.C. Thomas FThomas-at-NRCan.gc.ca, 902-426-4635, facsimile 902-426-6152 Natural Resources Canada, Bedford Institute of Oceanography, P.O. Box 1006, Dartmouth, Nova Scotia B2Y 4A2 Ressources naturelles Canada, l'Institut Oceanographique du Bedford, B.P. 1006, Dartmouth, (Nouvelle-Ecosse) B2Y 4A2 Government of Canada/Gouvernement du Canada
-----Original Message----- } From: dvanoekelen-at-omnilabo.be [mailto:dvanoekelen-at-omnilabo.be] Sent: Monday, March 15, 2004 7:07 PM To: microscopy-at-ns.microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dvanoekelen-at-omnilabo.be) from on Monday, March 15, 2004 at 01:30:08 ---------------------------------------------------------------------------
Email: dvanoekelen-at-omnilabo.be Name: Van Oekelen
Organization: Omnilabo
Title-Subject: [Microscopy] [Filtered] SEM from
Question: Dear Microscopists,
I know there are a lot of specialists regarding the use of SEM. Therefore, I was hoping that someone could help me finding a beautiful SEM image of a salt crystal.
Dear Frank, I have looked at lots of diamonds, both industrial in cutting tools and our hardness indenters. Some of them charge, but are otherwise unhurt by the beam. I know a geologist who uses one as a substrate for WDS analysis on small grains. Regards, Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 e-mail: mager-at-interchange.ubc.ca ----- Original Message ----- } From: "Thomas, Frank" {FThomas-at-nrcan.gc.ca} To: {microscopy-at-ns.microscopy.com} Sent: Tuesday, March 16, 2004 10:19 AM
Hello Listers:
I have a question regarding mouse brain hippocampus which normally does not contain any amount of IgG or IgM due to the blood/brain barrier. I am doing immunoelectron microscopy with a gold secondary which would be silver enhanced.
This experiment involves an injection/infection of a foreign protein sequence attached to amplicons to induce cells from CA1 region of the hippocampus to produce that protein. We see alot of cellular damage and the presence of inflammatory cells at the injection site along the needle tract. These lymphocytes and other cells involved in inflammation should produce IgG in the area of injury.
My question is: Would the IgG present in the inflammatory cells diffuse out into the neighboring CA1 to CA3 region? Could that cause a secondary antibody such as gold tagged goat anti-mouse F(ab')2 IgG or IgM to also label the surrounding normal cells(those which did not take up the foreign protein/amplicons) some distance from the injury? I am using a mouse monoclonal antibody to identify the infected cells which are producing this new protein.
Thanks for your comments!
Karen
Karen L. Bentley, M.S.(previously Jensen) Associate Scientist & Project Manager Electron Microscope Research Core University of Rochester Medical Center Rochester, NY 14642 585-275-1954
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 14:40:17 2004
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Your local JEOL service people should be able to assist. In the UK I think a good number of 120 kV and 200 kV TEMs have been converted.
Although I think SF6 is due to come under similar restrictions in 2-3 years. What happens then, I don't know.
I should declare an interest, as a JEOL employee. -- Larry Stoter NOTE - any message other than plain text will be automatically deleted :-)
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 14:40:18 2004
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Gallium FIB systems can certainly etch diamond (but then again, most things can be etched in a FIB). A few years ago DeBeers were looking at the possibility of using FIBs to put serial nos, etc on to gem quality diamonds - a pity nothing ever came of it (too easy to remove with a quick re-polish?) as SEM manufacturers were looking forward to jewellers having to buy SEMs to check out diamonds :-))
I would also suggest that in some newer thermal FEG SEMs, the electron probe current density can get very high, equivalent to terawatts per square metre, the sort of energy densities achieved in nuclear explosions. As has been pointed out, the thermal conductivity of diamond is very high but I wouldn't want to be the one to discover that this sort of energy density could damage gem quality diamonds :-)
I also recall many years ago looking at diamond in the TEM and that it was possible, with small probes to cause damage in electron-transparent diamond. This was thinned diamond crystals, as opposed to DLC or any other sort of film.
I should declare an interest, as an employee of JEOL, in selling SEMs and FIBs. -- Larry Stoter NOTE - any message other than plain text will be automatically deleted :-)
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 14:43:06 2004
} but there's no way that that an electron beam of say, } 15 or 20KeV can hurt a diamond in any way is there? ...
I am familiar with using diamond as a spectrum-free substrate for particle analysis at typical keV and m'probe beam currents (cathodo-luminescence being an aid over using polished vitreous carbon). They were commonly reused, and as far as I am aware we never damaged them in any way. Still, I believe you can feel 100% confident of using a defocused beam and lesser beam currents. The only caveat would be respect to any possible difference in altering the refractive index locally ... perhaps someone has possibly noticed(?)
I know that if you use the high voltage of a TEM and you have a poor vacuum with water vapor in it, you can drill holes in diamond when you focus the probe down. Don't know about SEM conditions.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472 Walck-at-PPG.com (412) 820-8651 (office) (412) 820-8515 (fax)
-----Original Message----- } From: Thomas, Frank [mailto:FThomas-at-nrcan.gc.ca] Sent: Tuesday, March 16, 2004 1:19 PM To: microscopy-at-ns.microscopy.com
This may be a stupid question, but there's no way that that an electron beam of say, 15 or 20KeV can hurt a diamond in any way is there? I was checking out a gemstone (unofficially - off the clock :-)for a colleague, which turned out to be a cubic zirconium, but then I got thinking - if it had been a diamond (carbon in crystal form)would that have been beam-sensitive in the way that organic (carbon-based) things are? My hunch is no, but if somebody ever asks me to check out their engagement ring to see if it's really diamond (yes, I hear people sometimes do that ;-), I don't want to be the guy that burns a hole in it.
Frank
F.C. Thomas FThomas-at-NRCan.gc.ca, 902-426-4635, facsimile 902-426-6152 Natural Resources Canada, Bedford Institute of Oceanography, P.O. Box 1006, Dartmouth, Nova Scotia B2Y 4A2 Ressources naturelles Canada, l'Institut Oceanographique du Bedford, B.P. 1006, Dartmouth, (Nouvelle-Ecosse) B2Y 4A2 Government of Canada/Gouvernement du Canada
-----Original Message----- } From: dvanoekelen-at-omnilabo.be [mailto:dvanoekelen-at-omnilabo.be] Sent: Monday, March 15, 2004 7:07 PM To: microscopy-at-ns.microscopy.com
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dvanoekelen-at-omnilabo.be) from on Monday, March 15, 2004 at 01:30:08 ---------------------------------------------------------------------------
Email: dvanoekelen-at-omnilabo.be Name: Van Oekelen
Organization: Omnilabo
Title-Subject: [Microscopy] [Filtered] SEM from
Question: Dear Microscopists,
I know there are a lot of specialists regarding the use of SEM. Therefore, I was hoping that someone could help me finding a beautiful SEM image of a salt crystal.
For some years, I have been imaging electron-dense inclusion bodies in various protozoal organisms by sticking them (unfixed, unsectioned) to formvar and using a relatively high voltage.
Lately I have been trying to do similar imaging in fixed specimens, because the fixation allows for the visualization of some membrane and vacuolar structure. Unfortunately, it also adds a lot of contrast--so much so that I often can't see the dense inclusions. I suspect this is because the the cells shrink a bit in the fixative medium, which renders them on the whole much more electron dense.
Any suggestions on alternative fixation procedures? I have tried various concentrations of glutaraldehyde and paraformaldehyde both alone and together with little luck. Or alternative TEM techniques? A colleague mentioned in passing, for instance, trying to find a scope with energy filtering capabilities, but I am not very familiar with this approach.
_______________________________ Peter Rohloff, PhD Laboratory of Molecular Parasitology Medical Scholars Program University of Illinois at Urbana-Champaign
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 15:24:35 2004
As a continuation of an earlier discussion last week; we are having a few problems getting the resin to infiltrate into the Bundle Sheath cells surronding vascular bundles in Ryegrass pseudostem.
We use an 812 resin and infiltrate overnight in 50/50 with Acetone and then 100% resin; two lots at 6-12 hours each. When we cut 1 um sections to view in LM looks OK; but then trim and polish the blockface for TEM sections we can sometimes see minute "holes" in the face where the resin has "fallen out" of the BS cells.
Any ideas on a possible solution to this annoying problem?
Thanks for your help
Raymond Bennett
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From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 15:51:10 2004
unless they have changed everything in immunology over the past year, the immunoglobulins are not going to diffuse out from the T cells, they will be actively secreted. i suspect you knew that, however, and are really interested in diffusion of antibodies within the surrounding tissues. lymphocytes will perform their programmed functions, wherever they happen to be. if they have crossed the blood brain barrier, or if they are actually astrocytes already resident, they will still perform their normal immunologic functions. you will get some interstitial antibody in the neighborhood of the lymphocytes, whether in the brain or any other tissue. if the foreign protein is to be trafficked to the plasmalemma, then you will have a potential problem telling whether the labelling is due to a reaction with primary mAb, or host produced polyclonal antibodies to the foreign protein. if the foreign protein is to be trafficked to the cell interior, then you should have less trouble, but you still may have to deal with the question of whether the reaction is due to the reaction of the primary mAb or host antibody which has been taken into the cell.
having said this, all need not be lost. have your investigators considered using human monoclonals produced in the Xenomouse model? the mouse is transgenic and produces human immunoglobulins, not murine. they could contact Abgenix about getting the antibody made. i do not think they would simply sell the mice for use as models, i think they would require you get the antibody from them or someone licensed by them to do the work. for that matter, i do not know whether they have actually licensed anyone, either.
no, i do not work for Abgenix, but i do know some people who wish to use their model, and we have discussed using direct and indirect immunogold EM with the project they wish to do.
Paul R. Hazelton, PhD Electron Microscope Unit University of Manitoba Department of Medical Microbiology 531 Basic Medical Sciences Building 730 William Avenue Winnipeg, Manitoba, Canada, R3E 0W3 e-mail: paul_hazelton-at-umanitoba.ca Phone:204-789-3313 Pager:204-931-954 Cell:204-781-1502 Fax:204-789-3926
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 16:47:42 2004
} For some years, I have been imaging electron-dense inclusion bodies in } various protozoal organisms by sticking them (unfixed, unsectioned) to } formvar and using a relatively high voltage. } } Lately I have been trying to do similar imaging in fixed specimens, } because the fixation allows for the visualization of some membrane and } vacuolar structure. Unfortunately, it also adds a lot of contrast--so } much so that I often can't see the dense inclusions. I suspect this is } because the the cells shrink a bit in the fixative medium, which } renders them on the whole much more electron dense. } } Any suggestions on alternative fixation procedures? I have tried } various concentrations of glutaraldehyde and paraformaldehyde both } alone and together with little luck. Or alternative TEM techniques? A } colleague mentioned in passing, for instance, trying to find a scope } with energy filtering capabilities, but I am not very familiar with } this approach. } Dear Peter, Do you have the facilities for cryo-fixation? Depending on the thickness(es) of the organisms you might have success either with plunge-freezing--for ~ {1 um--or high-pressure-freezing followed by cryo-substitution and sectioning or by cryo-sectioning. Energy-filtered EM could be done either by taking a conventional image, then acquiring spectra from areas of interest, or--probably more usefully in your case--by acquiring an image using only elastically scattered electrons (a zero-loss image) or electrons that have lost energy in a window characteristic of a particular element (element mapping). The first method will enhance contrast due to atoms heavier than those predominant in biological molecules, and the second method will show you where the amount of a particular element is high. A disadvantage of element mapping is that it requires a high electron dose, since electrons losing energies characteristic of a particular element are a small fraction of the transmitted electrons. Energy filtering is especially useful if the inclusions consist of element(s) not present in large concentration(s) in the rest of the organism. AFAIK, imaging filters are available only for scopes with HV {= 400 kV (which may or may not include "relatively high voltage"). Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 16:53:28 2004
Cell walls can sometimes be difficult to infiltrate. We usually dehydrate with propylene oxide after our last 100% ETOH and then use ~33% steps of resin:PO mixtures to infiltrate. A couple of hours in 33% resin:66% PO, 4 hrs.to overnight in 66% resin : 33% PO, and the two or three changes of 100% resin, the first overnight and the next two 6 - 12hrs. each. All infiltration steps are carried out on a sample rotator. I do not remember, but do the BS cells have silica bodies? If so the holes that you are seeing may be the silica bodies pulling out of the block face. In this case you are on your own!
Raymond Bennett wrote:
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-- ================================================================== Greg Strout Electron Microscopist, University of Oklahoma WWW Virtual Library for Microscopy: http://www.ou.edu/research/electron/www-vl/ e-mail: gstrout-at-ou.edu (405)325-4391 Opinions expressed herein are mine and not necessarily those of the University of Oklahoma ==================================================================
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 16 18:57:10 2004
Dear Karen I am sorry if I don't understand you right. Many cells have Ig-like receptors on their membranes including macrophages (I do believe). Brain itself has a cells of macrophagal origin (glial cells for instance) and your injury broke blood barrier, so more cells migrate to the brain (mostly macrophages). At such background, it seems to me, you may not use anti-mouse Ig-s as a secondary AB. It's common practice to use, for instance, rabbit primary ABs and anti-rabbit secondary. You also need to do negative control with secondary ABs only. Forgive me if I did not understand your question. Sergey.
At 12:52 PM 3/16/2004, you wrote:
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_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry 10833 Le Conte Ave, Room 33-089 Los Angeles, CA 90095
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (nessonm-at-onid.orst.edu) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Tuesday, March 16, 2004 at 19:30:27 ---------------------------------------------------------------------------
Email: nessonm-at-onid.orst.edu Name: Michael Nesson
Organization: Oregon State University
Title-Subject: [Microscopy] [Filtered] MListserver: SiO grid films
Question: Can anyone provide a protocol for making SiO filmed gold grids? I'm planning some oxygen plasma-ashing experiments on some amorphous metal-rich biological materials. I know that I can obtain such grids from several of the EM supply houses, but I don't want to pay the prices, if I can make them myself. I have available an oil-diffusion pumped evaporator, tungsten baskets, and SiO granules.
The last part of the letter listed below indicates damage to Nikon lenses by the use of Zeiss immersion oil. Do you, Ralph, or anyone else have instances where this has occurred? (I would suspect that this would occur on inverted scopes most often and would be the result of dissolving the seal between the lens and the outer case.) If so was this the old Zeiss or the new Zeiss oil immersion medium? I've got Nikon scopes and would like to post messages in the appropriate rooms if this has proven to be a problem. I also have Zeiss oil over 15 years old with no solidification problems that I've kept hidden.
Also, does anyone know where to find small plastic 5-10 ml bottles that would be suitable for dispensing oil immersion medium?
.. Jerry Calvin
listed 3-3-04 .. Also be aware that different types of immersion oil should never be allowed to mix, and that some types of oil can damage the mounting cement of some brands of lenses. Using Zeiss immersion oil with some Nikon lenses, for example, can be disastrous.
Ralph Common Electron Microscopist Michigan State University Division of Human Pathology A608 East Fee Hall East Lansing, MI 48824 517-355-7558; fax 517-432-1053 ralph.common-at-ht.msu.edu -- **************************** Jerry G. Calvin Science Support technician Box 0731 Biology Department Vassar College 124 Raymond Avenue Poughkeepsie, NY 12604-0731
Second and Final Call for Papers Michigan Microscopy &Microanalysis Society Spring Meeting in 2004
Bavarian Inn Frankenmuth, MI April 16, 2004
Revised Abstract Deadline: March 26, 2004
The Spring Meeting of the Michigan Microscopy and Microanalysis Society will be held on Friday April 16, 2004 at Bavarian Inn in Frankenmuth, MI. In this one-day conference, there will be two sessions. One is a platform session. This session will have approximately 8-10 speakers representing industry, academia, and research laboratories. The other one is a poster session newly opened this year. Please encourage your colleagues who prefer to avoid a platform presentation to submit abstracts for the poster presentations. In addition to the speakers, vendors will exhibit a wide range of products and services of interest to the microscopy community. Presentations are being solicited from researchers in the Physical and Biological Sciences, including one vendor presentation and two invited speakers. Student participation is particularly encouraged. Also, vendors are encouraged to contact the below address to reserve space for product display.
Abstract Submission Please submit a 300 to 350 word abstract by March 26th indicating which session you prefer to:
Ginam Kim (Ph.D) or email to g.kim-at-dowcorning.com Dow Corning Corp. P.O. Box 994, C041D1 Midland, MI 48686 Tel) (989)-496-5077 Fax) (989)-496-5121
The poster format information will be provided to participants at a later date.
Kevin Battjes Impact Analytical Voice 989-832-5555, ext 556 Michigan Molecular Institute Fax 989-832-5560 1910 W. St Andrews Road e-mail: battjes-at-mmi.org Midland MI 48640 battjes-at-impactanalytical.com
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 17 12:31:19 2004
Greetings, In connection to the thread, at one time in the past different immersion oils from the different 'scope makers had diferent refractive indices. Therefore, if they got mixed, the results were really bad, and presumably the lens would have been designed around a given index. I *think* that the oils from Zeiss, Nikon, Olympus, Leica now have the same refractive index, but I am not sure, and I would be interested to hear if anyone knows.
Tobias Baskin } } } The last part of the letter listed below indicates damage to Nikon } lenses by the use of Zeiss immersion oil. Do you, Ralph, or anyone } else have instances where this has occurred? (I would suspect that } this would occur on inverted scopes most often and would be the } result of dissolving the seal between the lens and the outer case.) } If so was this the old Zeiss or the new Zeiss oil immersion medium? } I've got Nikon scopes and would like to post messages in the } appropriate rooms if this has proven to be a problem. I also have } Zeiss oil over 15 years old with no solidification problems that } I've kept hidden. } } Also, does anyone know where to find small plastic 5-10 ml bottles } that would be suitable for dispensing oil immersion medium? } } .. Jerry Calvin } } listed 3-3-04 } .. Also be aware that different types of immersion oil should never } be allowed to mix, and that some types of oil can damage the } mounting cement of some brands of lenses. Using Zeiss immersion oil } with some Nikon lenses, for example, can be disastrous.
Some "oils" are not oils at all, at least not in the sense that they are derived from petroleum, or contain fully saturated long-carbon chain (aliphatics/alkanes, etc) compounds. Virtually any substance that has the correct RI, with useable physical properties will be suitable as an immersion "oil". Unfortunately, this leaves the door wide open for manufacturers to use anything from a "real" oil, to compounds like benzyl benzoate, or mixtures of a liquid with a dissolved solid component. The latter case is probably what has caused the infamous Zeiss crystallizing-oil problem. The Zeiss oil is probably saturated when it comes from the factory. When it experiences a drop in temperature, or a bit of evaporation, the solution becomes supersaturated and the crystals drop out. As has been mentioned, warming the mix redissolves the crystals. At that point, I would imagine the RI of the oil would be too high, and would create spherical aberration.
In any case, if you mix oils on the same microscope slide or objective lens without thorough cleaning, chances are you'll create a blurry liquid-liquid interface, and there goes the resolution! Since all the microscope manufacturers know that their oil might not be miscible with any other manufacturers oil, they always recommend that you pick one oil and use it exclusively.
Jim
-----Original Message----- } From: Tobias Baskin [mailto:baskin-at-bio.umass.edu] Sent: Wednesday, March 17, 2004 1:50 PM To: microscopy-at-MSA.microscopy.com
Greetings, In connection to the thread, at one time in the past different immersion oils from the different 'scope makers had diferent refractive indices. Therefore, if they got mixed, the results were really bad, and presumably the lens would have been designed around a given index. I *think* that the oils from Zeiss, Nikon, Olympus, Leica now have the same refractive index, but I am not sure, and I would be interested to hear if anyone knows.
Tobias Baskin } } } The last part of the letter listed below indicates damage to Nikon } lenses by the use of Zeiss immersion oil. Do you, Ralph, or anyone } else have instances where this has occurred? (I would suspect that } this would occur on inverted scopes most often and would be the } result of dissolving the seal between the lens and the outer case.) } If so was this the old Zeiss or the new Zeiss oil immersion medium? } I've got Nikon scopes and would like to post messages in the } appropriate rooms if this has proven to be a problem. I also have } Zeiss oil over 15 years old with no solidification problems that } I've kept hidden. } } Also, does anyone know where to find small plastic 5-10 ml bottles } that would be suitable for dispensing oil immersion medium? } } .. Jerry Calvin } } listed 3-3-04 } .. Also be aware that different types of immersion oil should never } be allowed to mix, and that some types of oil can damage the } mounting cement of some brands of lenses. Using Zeiss immersion oil } with some Nikon lenses, for example, can be disastrous.
First of all, I'd like to thank all of those helpful people who commented on the histo-diamond knives by Diatome. I am very impressed with what I heard and I will purchase one as soon as possible.
Still, I am wondering though how most people who use this knife for semi-thin (0.5 microns)sectioning rough in their specimens. Do you people still use glass knives, or do you just go very slowly with the histoknife. I thought that it might be prudent to still use glass knives for this purpose, if only to spare the histo diamond knife from rough treatment.
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 17 16:00:32 2004
I trim with a blade and then rough the block face directly with the histo knife. Works great.
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: Garry Burgess [mailto:GBurgess-at-exchange.hsc.mb.ca] Sent: Wednesday, March 17, 2004 5:03 PM To: Microscopy-at-sparc5.microscopy.com
First of all, I'd like to thank all of those helpful people who commented on the histo-diamond knives by Diatome. I am very impressed with what I heard and I will purchase one as soon as possible.
Still, I am wondering though how most people who use this knife for semi-thin (0.5 microns)sectioning rough in their specimens. Do you people still use glass knives, or do you just go very slowly with the histoknife. I thought that it might be prudent to still use glass knives for this purpose, if only to spare the histo diamond knife from rough treatment.
From MicroscopyL-request-at-ns.microscopy.com Wed Mar 17 17:52:22 2004
On Mar 17, 2004, at 4:11 PM, Alexander Cronin wrote:
Dear Alex,
} 1. What is the most efficient material for electron diffraction? (I } want } high flux in one diffraction order).
Since the scattering cross section increases with increasing Z, I'd guess that the most efficient material reasonably available would be uranium metal; however, the most practical efficient material is likely to be gold (or maybe bismuth). In addition to the scattering cross section, the crystal form would be important to give high intensity in a single diffracted spot, so someone who knows more about crystallography than I do can correct me if I'm off base. } } 2. Has anybody seen electron diffraction from fabricated gratings?
The wavelength of an electron typically used in TEM is very short, so diffraction from a grating produced by making grooves on a substrate is not likely to work, since the variations in groove spacings will be larger than the wavelength; however, materials grown epitaxially, if you consider them to be gratings, do show ED, and maybe low-angle reflection ED has been seen from more conventional gratings--once again, someone more expert than I should comment. } } } 3. How about evidence that diffracted electron-waves are phase } coherent } with the incident beam?
I think that can be determined from holography; I'll let the experts comment. The incident beam itself varies in coherence depending on the source; a W filament is the least coherent of the usual sources, a LaB6 filament is more coherent and a field emission gun is the most coherent. (There may be even more coherent non-conventional sources.) } } 4. I seek references for building a three-grating (Mach-Zhender style) } electron interferometer to study the Aharonov-Bohm effect.
Good luck. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 08:12:29 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (verlaj-at-medicine.ufl.edu) from on Wednesday, March 17, 2004 at 12:38:29 ---------------------------------------------------------------------------
Organization: University of Florida College of Medicine
Title-Subject: [Microscopy] [Filtered] MListserver: seeking recommendations for a new carbon coater
Question: Dear Microscopy Listservers, We are strongly considering replacing our old Denton evaporator with a new unit to carbon coat grids. I would be grateful for the opinions of anyone who has purchased a new unit in the recent past. I am particularly interested in the reliability of the unit and the quality of the coating. We are using the Denton almost exclusively to coat Formvar-coated grids for supporting and stabilizing Lowicryl thin sections. Ease of use and cost are other factors we will consider. Vendors are more than welcome to contact me directly.
Thanks in advance for your time and the benefits of your experience.
Jill
Jill Verlander Reed, DVM Director, College of Medicine Electron Microscopy Core Facility University of Florida P.O. Box 100215 Health Science Center 1600 SW Archer Road Room RB-167 Gainesville, FL 32610-0215 telephone (352) 846-0820 facsimile (352) 392-8996
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (ptibbits-at-emerson-ept.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, March 17, 2004 at 12:19:44 ---------------------------------------------------------------------------
Email: ptibbits-at-emerson-ept.com Name: Patrick Tibbits
Organization: Emerson Power Transmission
Title-Subject: [Microscopy] [Filtered] MListserver: SEM Maintenance
Question: I would like to get suggestions for companies which carry maintenance contracts.
Also, I would like to learn of any third-party (not OEM) maintenance providers.
I have a Hitachi 2460N SEM and Oxford 300 ISIS EDS.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (hadden-at-wingate.edu) from http://microscopy.com/MicroscopyListserver/MLFormMail.html on Wednesday, March 17, 2004 at 13:50:41 ---------------------------------------------------------------------------
Question: One of my SEM students is working with teeth. Can anyone recommend the best way to view the teeth in the SEM, i.e. hi vac SEI or low vac BEI [or any other suggestions for most effective imaging of enamel structure]? We are using a JEOL LV5600 SEM.
Thanks in advance.
Lee Hadden Department of Biology Wingate University Wingate, NC 28174
I have a question about lightsources in microscopy. For fluorescence microscopy you can use Mercury or Xenon arcs and I am curious about the level of even illumination you can achieve with this kind of lightsources. If they would be pointsources the "radial" intensity would diminish with 1/radius^3, but this doesn't seem to be the case ?
What is the residual uneven illumination below which it is not possible to get with a traditional Xenon or Mercury Arc style illumination ?
Long ago I once read that there are fluorescent samples which you coud use to make a background profiles for fluorescetce microscopy ? are these pieces of plastic in which a fluorochorem is embedded ?
Has anyone ever measured the contribution of multiwell plate bottoms to the illumination profile, due to the non-flat bottoms acting as a lens ?
Regards,
Peter Van Osta
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 12:37:09 2004
On Mar 18, 2004, at 6:32 AM, Jill Verlander Reed wrote:
} We are strongly considering replacing our old Denton evaporator } with a new unit to carbon coat grids. } I would be grateful for the opinions of anyone who has purchased } a new unit in the recent past. } I am particularly interested in the reliability of the unit and } the quality of the coating. We are using the Denton almost } exclusively to coat Formvar-coated grids for supporting and } stabilizing Lowicryl thin sections. Ease of use and cost are other } factors we will consider. } Vendors are more than welcome to contact me directly. } } Thanks in advance for your time and the benefits of your experience. } Dear Jill, About a year ago we purchased a Cressington 208 evaporator, which has a turbopump, to do both carbon and metal evaporation. The unit has been completely reliable, and the coatings have been good. The best coatings are obtained when the vacuum is around 10^-5, which takes a fairly long time to achieve (depending on how much filter paper, etc., in put into the chamber). The instrument is very easy to use for carbon evaporation, and slightly less so for metal evaporation when the metal piece has to be loaded in a wire basket, and the cost was very reasonable for a turbopump desktop system--~$20k if you only want the carbon power supply, ~$25k with both carbon and metal power supplies. Yours, Bill Tivol, PhD EM Scientist and Manager Cryo-Electron Microscopy Facility Broad Center, Mail Code 114-96 California Institute of Technology Pasadena CA 91125 (626) 395-8833 tivol-at-caltech.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 12:52:22 2004
We have projects doing teeth and bone here. Since they are not generally interested in soft tissues or cells, we just air dry the teeth/bone, sputter coat and go look. Works well. This is in a Hitachi S-570 with LaB6, or our S-900 FESEM, but I've also done this in conventional tungsten-filament SEMs. High vacuum, SEI and BSE, whole teeth or broken. I've got a dinosaur tooth project that should be coming in, and he's likely to do some acid-etching of the teeth to bring out the enamel, but that's the only preparation step. There's also this site from the American Museum of Natural History: http://research.amnh.org/vertpaleo/enamel/index.html It's on preparing tooth enamel for SEM.
Phil
} Email: hadden-at-wingate.edu } Name: Lee Hadden } } Organization: Wingate University } } Title-Subject: [Microscopy] [Filtered] MListserver: } } Question: One of my SEM students is working with teeth. Can anyone } recommend the best way to view the teeth in the SEM, i.e. hi vac SEI } or low vac BEI [or any other suggestions for most effective imaging } of enamel structure]? We are using a JEOL LV5600 SEM. } } Thanks in advance. } } Lee Hadden } Department of Biology } Wingate University } Wingate, NC 28174 } } ---------------------------------------------------------------------------
-- Philip Oshel Supervisor, BBPIC microscopy facility Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 - 1284 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 13:42:02 2004
I'm looking for a recipe or protocol for Fekete's Fixative.
please help!
Thanks
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From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 14:01:44 2004
} Larry; } } Just curious but doesn't Debeers laser mark their gems? I saw a } program on TV about synthetic diamonds, gem quality, and they stated } the difficulty in identification for fraud and authenticity, e.g. } real or man-made. } } Is this a fact? I would imagine the laser marking could be polished off? } } Peter Tomic } I also seem to recall something on laser marking. As with FIB marking, I guess is could be easily polished off. Although, if you have a valuable gem-quality diamond with lots of facets, that may be a much bigger job than most people would want to undertake, especially as you would not want to devalue the stone.
I also seem to remember reading somewhere about a method for using two laser beams - each on its own had no effect but where they crossed, there was sufficient energy to damage the diamond. This then makes it possible to put the serial number inside the diamond, so it can't be polished out without seriously reducing the value. -- Larry Stoter NOTE - any message other than plain text will be automatically deleted :-)
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 14:36:15 2004
Greetings, I'm trying to find a good cocktail for non-specific digestion of tissue from bone samples--only the mineralized bone needs to be left unscathed. I'm looking for a relatively cheap solution that will yield nice clean bones. I'm aware of a product from Fisher, but it costs about $70/100ml. It seems to me that there must be a simple solution that I'm just not aware of (aside from culturing maggots). Thanks in advance for any suggestions. Regards, Karl G.
-- Karl Garsha Light Microscopy Specialist Imaging Technology Group Beckman Institute for Advanced Science and Technology University of Illinois at Urbana-Champaign 405 North Mathews Avenue Urbana, IL 61801 Office: B650J Phone: 217.244.6292 Fax: 217.244.6219 Mobile: 217.390.1874 www.itg.uiuc.edu
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 14:51:54 2004
Household bleach, 1 part bleach to 4 parts water. or Adolph's meat tenderizer is papain, don't know the concentration needed. You can get papain form Sigma, works well at 37C. I don't know any other details.
Karl Garsha wrote:
} Greetings, } I'm trying to find a good cocktail for non-specific digestion of } tissue from bone samples--only the mineralized bone needs to be left } unscathed. I'm looking for a relatively cheap solution that will } yield nice clean bones. I'm aware of a product from Fisher, but it } costs about $70/100ml. It seems to me that there must be a simple } solution that I'm just not aware of (aside from culturing maggots). } Thanks in advance for any suggestions. } Regards, } Karl G. }
-- -- ********************************************** Geoff McAuliffe, Ph.D. Neuroscience and Cell Biology Robert Wood Johnson Medical School 675 Hoes Lane, Piscataway, NJ 08854 voice: (732)-235-4583; fax: -4029 mcauliff-at-umdnj.edu **********************************************
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 15:23:20 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (heather.eberhardt-at-frx.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Thursday, March 18, 2004 at 08:34:33 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] Thin sectioning of tablets
Question: Hello everyone, Iím new here but learning a lot. I need to take thin sections of pharmaceutical tablets, and need several sections from a single tablet (Ideally 10+). They are for use with optical microscopy, ideally PLM. I am trying to figure out which type of epoxy compound I should be using since there are so many out there. Any help or suggestions from someone who has experience in this would be appreciated.
Thanks,
Heather Eberhardt
Heather Eberhardt Materials Characterization Group Forest Research Institute Hauppauge, NY 631-436-2619
} I'm looking for a recipe or protocol for Fekete's Fixative. } } please help! } } Thanks
********** Twice in one week? Wow. Here are some ref's: Amer. Coll. of Vet Path. 2003 meeting abstract Poster T-6, Abst. 132 Toxocol. Path). Its an acid alcohol-formalin fix. See also: Am J Physiol Gastrointest Liver Physiol 274: G544-G551, 1998; 0193-1857/98 $5.00 Vol. 274, Issue 3, G544-G551, March 1998
Differential susceptibility of inbred mouse strains to dextran sulfate sodium-induced colitis
Michael Mähler1, Ian J. Bristol1, Edward H. Leiter1, Aletha E. Workman1, Edward H. Birkenmeier,1, Charles O. Elson2, and John P. Sundberg1
-- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Thu Mar 18 16:17:14 2004
WE DO IT ALL , WE ARE REASONABLE AND WE ARE COMPETENT. PLEASE CALL IF INTERESTED. WE ARE NOT TELEMARKETER TYPES, SO WE DO NOT INUNDATE YOU WITH ACCOLADES OF OUR SELVES.
IMAQUE IMAGING BEN GHAFFARI 703-379-0027 571-437-6593 703-257-6321 FAX ----- Original Message ----- } From: "by way of MicroscopyListserver" {ptibbits-at-emerson-ept.com} To: {microscopy-at-ns.microscopy.com} Sent: Thursday, March 18, 2004 9:31 AM
Dear Larry, I believe the diamonds mined in Canada are all marked with a laser-etched symbol, some with a polar bear and some with a stylized maple leaf, to identify them as non-conflict diamonds. They are advertised as such. Regards, Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 e-mail: mager-at-interchange.ubc.ca ----- Original Message ----- } From: "Larry Stoter" {larry-at-cymru.freewire.co.uk} To: "Tomic, Peter (Peter)" {ptomic-at-agere.com} ; {Microscopy-at-MSA.Microscopy.Com} Sent: Thursday, March 18, 2004 9:56 AM
Quick internet search found the following which appears to be good for eyes.
Fekete's acid-alcohol-formalin fixative (3.2% formaldehyde, 0.7 M acetic acid, 61% ethanol). After 24 h, the fixative was replaced with 70% ethanol. Eyes were embedded in paraffin, sectioned (5 micron) and stained with haematoxylin and eosin (H&E) using standard procedures.
Gill Brown
Histopathology Group Asthma and Allergy Disease Biology ri- CEDD. GlaxoSmithKline Medicines Research Centre,
----- Forwarded by Gillian 2 Brown/PharmRD/GSK on 19-Mar-2004 08:39 -----
"Louro, Pedro" {pedro.louro-at-spcorp.com}
18-Mar-2004 20:00
To: Microscopy
cc: Subject: [Microscopy] Fekete's Fixative
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I'm looking for a recipe or protocol for Fekete's Fixative.
please help!
Thanks
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From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 08:53:24 2004
Hello Peter, Getting truly even illumination out of a conventional fluorescence arc lamp isn't a trivial task. The illumination source doesn't really approximate a point source very well--in a conventional lamp housing one can adjust the position of the light bulb and mirror so there is an image of the arc and a reflected image of the arc coming from behind, this is defocused to provide quasi even illumination with the intensity highest in the center of the field of view. Depending on the centration of the objective and filters, the peak intensity can wander. The most even illumination I've seen from an arc lamp is achieved by fiber-coupling the lamp output to a fiber optic bent around a large radius to homogenize the light-the fiber output provides a better approximation of a point source and the output at the end of the fiber is even. The Applied Precision Delta Vision takes this approach. It isn't an easy task to couple an arc lamp to a fiber optic, though. There are fluorescent plastic slides --I believe they can be obtained from Chroma. On an inverted microscope, one can use dilution series of fluorochromes to test things; I prefer this approach (I use chambered coverslips from LabTek to hold the fresh solutions of fluorochrome). I haven't heard of anyone measuring the optical abberations caused by multiwell plates, but it would seem that image arithmetic could be used to quantify the difference in fluorescence intensity across the field of view using such plates as opposed to using chambered coverslips with a flat .17mm borosilicate glass bottom. The fluorescent dilutions would be helpful in this context. Regards, Karl
Peter Van Osta wrote:
} } } ------------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } } Hi, } } I have a question about lightsources in microscopy. For fluorescence } microscopy you can use Mercury or Xenon arcs and I am curious about } the level of even illumination you can achieve with this kind of } lightsources. If they would be pointsources the "radial" intensity } would diminish with 1/radius^3, but this doesn't seem to be the case ? } } What is the residual uneven illumination below which it is not } possible to get with a traditional Xenon or Mercury Arc style } illumination ? } } Long ago I once read that there are fluorescent samples which you coud } use to make a background profiles for fluorescetce microscopy ? are } these pieces of plastic in which a fluorochorem is embedded ? } } Has anyone ever measured the contribution of multiwell plate bottoms } to the illumination profile, due to the non-flat bottoms acting as a } lens ? } } Regards, } } Peter Van Osta } }
-- Karl Garsha Light Microscopy Specialist Imaging Technology Group Beckman Institute for Advanced Science and Technology University of Illinois at Urbana-Champaign 405 North Mathews Avenue Urbana, IL 61801 Office: B650J Phone: 217.244.6292 Fax: 217.244.6219 Mobile: 217.390.1874 www.itg.uiuc.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 09:28:31 2004
I'm evaluating backscatter detectors, and am in need of this familiar test material ... i.e., a difference in Z of 0.1 at Z=30. I see references to this standard in the supplies catalogs, but it is not immediately available, and the picture of it implies that it's not even polished ... which would contribute a lot to broadening the image's histogram peaks. It would help a lot if I could find a another source, and polish a sample of it next week. Is anyone aware of a poor mans' source of inclusion-free alpha-beta (or duplex) brass?
I don't even suppose it needs to be specifically AB brass, because all I need do is polish it extremely well, that it have neighboring phases to contrast, and have an average Z } 22 ... and make comparisons with the same material. For example, I have "pure" FeS exsolved in pyrrhotite, but the scale doesn't really allow me to select a relatively large area of each for determining the noise.
Suggestions most welcome ..
tia & cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 09:41:43 2004
Quoting "Louro, Pedro" {pedro.louro-at-spcorp.com} : --------------------------------------------------------------- } I'm looking for a recipe or protocol for Fekete's Fixative
Pedro,
I believe that the original regerence is in AM. J. Path. 14:557, 1938.
This reference I found in Histopathologic Technic and Practical Histochemistry by R.D. Lillie, 1954 in which the formula is stated as 10 cc. 37-40% formaldehyde 5 cc. Glacial acetic acid 100 cc. 70% alcohol
It states that it is a good glycogen preservative and suggests leaving the tissue in 70% alcohol rather than in the fixative if it is necessary to store the tissue for any length of time after fixation.
I love reading old histology books! Pat Connelly psconnel-at-sas.upenn.edu Research Specialist Dept. of Biology Univ. of Pennsylvania Philadelphia, PA 19104
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 10:13:19 2004
Hi Peter, i totally agree with Karl.... the fibercoupling is a way, for certain (blue) wavelengths you want to use liquid wave guides - but you will get a new problem once you bring your fibers/LWG`s in a different position. so you have definitively to fixate them somewhere on your scope! maybe you could make a "flatness" around 10%! if that`s not enough, you will have to correct the images anyways. just in case: correctedimage=(rawimage-rawdark)/(flatimage-flatdark) ["dark" are the darkcounts with ccd... the "dark" image you get with light turned off] propably you want to make a new flatfield image from time to time, since the lamp changes the properties in terms of resulting flatness incredibly over the time! we did at least once a week with our (Zeiss) HBO100. i wouldn`t suggest using bulk plastic slides, since these will homogenize your illumination quite a bit. you will see that clearly if you try to stitch the resulting images together to create a bigger image. The way Karl is describing seems much better, especially because you would be able to control the resulting intensity by concentration. we had the best results with a thin (1..3µm) fluorescent layer e.g. fluorophore-doped
polyimide on borosilicate-glass for flatfielding. its the same idea, just in a so.lid
phase. these are just easier to handle, store and use as dilutions... especially if you want to use the same thing different times.
cheers thomas
Karl Garsha schrieb:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } Hello Peter, } Getting truly even illumination out of a conventional fluorescence arc } lamp isn't a trivial task. The illumination source doesn't really } approximate a point source very well--in a conventional lamp housing one } can adjust the position of the light bulb and mirror so there is an } image of the arc and a reflected image of the arc coming from behind, } this is defocused to provide quasi even illumination with the intensity } highest in the center of the field of view. Depending on the centration } of the objective and filters, the peak intensity can wander. } The most even illumination I've seen from an arc lamp is achieved by } fiber-coupling the lamp output to a fiber optic bent around a large } radius to homogenize the light-the fiber output provides a better } approximation of a point source and the output at the end of the fiber } is even. The Applied Precision Delta Vision takes this approach. It } isn't an easy task to couple an arc lamp to a fiber optic, though. } There are fluorescent plastic slides --I believe they can be } obtained from Chroma. On an inverted microscope, one can use dilution } series of fluorochromes to test things; I prefer this approach (I use } chambered coverslips from LabTek to hold the fresh solutions of } fluorochrome). } I haven't heard of anyone measuring the optical abberations } caused by multiwell plates, but it would seem that image arithmetic } could be used to quantify the difference in fluorescence intensity } across the field of view using such plates as opposed to using chambered } coverslips with a flat .17mm borosilicate glass bottom. The fluorescent } dilutions would be helpful in this context. } Regards, } Karl } } Peter Van Osta wrote: } } } } } } } ------------------------------------------------------------------------------ } } } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- } } http://www.msa.microscopy.com/MicroscopyListserver } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } ------------------------------------------------------------------------------- } } } } } } Hi, } } } } I have a question about lightsources in microscopy. For fluorescence } } microscopy you can use Mercury or Xenon arcs and I am curious about } } the level of even illumination you can achieve with this kind of } } lightsources. If they would be pointsources the "radial" intensity } } would diminish with 1/radius^3, but this doesn't seem to be the case ? } } } } What is the residual uneven illumination below which it is not } } possible to get with a traditional Xenon or Mercury Arc style } } illumination ? } } } } Long ago I once read that there are fluorescent samples which you coud } } use to make a background profiles for fluorescetce microscopy ? are } } these pieces of plastic in which a fluorochorem is embedded ? } } } } Has anyone ever measured the contribution of multiwell plate bottoms } } to the illumination profile, due to the non-flat bottoms acting as a } } lens ? } } } } Regards, } } } } Peter Van Osta } } } } } } -- } Karl Garsha } Light Microscopy Specialist } Imaging Technology Group } Beckman Institute for Advanced Science and Technology } University of Illinois at Urbana-Champaign } 405 North Mathews Avenue } Urbana, IL 61801 } Office: B650J } Phone: 217.244.6292 } Fax: 217.244.6219 } Mobile: 217.390.1874 } www.itg.uiuc.edu
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 10:33:46 2004
We have a new SEM/EDS system at NSAC. This is the first electron microscope in our College. The College administration has asked me to develop a policy for usage, funding, prioritization and data collection of the SEM/EDS unit. The administration could not provide a technical support for the unit. I will be the major user of the system, I would assume there will be between 10 and 15 other users from the College. The system has been purchased with CFI funding to support research programs.
I would appreciate your comments and suggestions on how to develop user fees, how much of my time should be dedicated to the unit as teaching others, managing the lab, and simple maintenance. Perhaps I can ask the administration that 30-40% of my time to be dedicated to the unit..
Kind regards Valtcho Jeliazkov
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 12:05:16 2004
Although I am not in a university environment, I would strongly urge you to develop a thorough training program with specific instructions on how not to break things. Where people get in trouble, in my experience, is in areas like crashing the stage into the EDX nosepiece and lens, as well as putting in samples that are not thoroughly dry, [except for ESEMS], sample exchange and/or inappropriate samples for vacuum environments. The more you have written down in terms of instruction, the less likely it will be that your 30-40% of SEM time will be with wrench in hand. This will also help when you need to exile someone from the lab. since you will have documented procedures to point to. It also impresses upper management. In your case upper faculty and admins.
With regard to EDX, make them calibrate before using. It's a good habit to have.
Hope this is of some value.
Peter Tomic Agere Systems
-----Original Message----- } From: Valtcho Jeliazkov (Zheljazkov) Ph.D. [mailto:vjeliazkov-at-nsac.ns.ca] Sent: Friday, March 19, 2004 11:52 AM To: Microscopy-at-MSA.Microscopy.Com
Hi all,
We have a new SEM/EDS system at NSAC. This is the first electron microscope in our College. The College administration has asked me to develop a policy for usage, funding, prioritization and data collection of the SEM/EDS unit. The administration could not provide a technical support for the unit. I will be the major user of the system, I would assume there will be between 10 and 15 other users from the College. The system has been purchased with CFI funding to support research programs.
I would appreciate your comments and suggestions on how to develop user fees, how much of my time should be dedicated to the unit as teaching others, managing the lab, and simple maintenance. Perhaps I can ask the administration that 30-40% of my time to be dedicated to the unit..
Kind regards Valtcho Jeliazkov
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 12:27:31 2004
Do you have a maintenance contract ? The policy specifics and your time allotment may hinge on this factor.
Regards, Ed
----- Original Message ----- } From: "Valtcho Jeliazkov (Zheljazkov) Ph.D." {vjeliazkov-at-nsac.ns.ca} To: {Microscopy-at-MSA.Microscopy.Com} Sent: Friday, March 19, 2004 8:52 AM
PS to Peters comments -
Very Important also..keep an ongoing log near the system to record all maintenance and 'problems'...this is invaluable for troubleshooting thoughout the life of the system.
Also, in the training process, cover the 'artifact' issues (SEM as well as EDS). Never rely on the automation!
} From: "Tomic, Peter (Peter)" {ptomic-at-agere.com} } Date: Fri, 19 Mar 2004 13:23:39 -0500 } To: "Valtcho Jeliazkov (Zheljazkov) Ph.D." {vjeliazkov-at-nsac.ns.ca} , } {Microscopy-at-MSA.Microscopy.Com} } Subject: [Microscopy] RE: SEM/EDS system policy for usage } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- --} - } } Valtcho; } } Although I am not in a university environment, I would strongly urge you to } develop a thorough training program with specific instructions on how not to } break things. Where people get in trouble, in my experience, is in areas like } crashing the stage into the EDX nosepiece and lens, as well as putting in } samples that are not thoroughly dry, [except for ESEMS], sample exchange } and/or inappropriate samples for vacuum environments. The more you have } written down in terms of instruction, the less likely it will be that your } 30-40% of SEM time will be with wrench in hand. This will also help when you } need to exile someone from the lab. since you will have documented procedures } to point to. It also impresses upper management. In your case upper faculty } and admins. } } With regard to EDX, make them calibrate before using. It's a good habit to } have. } } Hope this is of some value. } } Peter Tomic } Agere Systems } } -----Original Message----- } } From: Valtcho Jeliazkov (Zheljazkov) Ph.D. } [mailto:vjeliazkov-at-nsac.ns.ca] } Sent: Friday, March 19, 2004 11:52 AM } To: Microscopy-at-MSA.Microscopy.Com } Subject: [Microscopy] SEM/EDS system policy for usage } } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- --} - } } Hi all, } } We have a new SEM/EDS system at NSAC. This is the } first electron microscope in our College. The } College administration has asked me to develop a } policy for usage, funding, prioritization and data } collection of the SEM/EDS unit. The } administration could not provide a technical } support for the unit. } I will be the major user of the system, I would } assume there will be between 10 and 15 other users } from the College. The system has been purchased } with CFI funding to support research programs. } } I would appreciate your comments and suggestions } on how to develop user fees, how much of my time } should be dedicated to the unit as teaching } others, managing the lab, and simple maintenance. } Perhaps I can ask the administration that 30-40% } of my time to be dedicated to the unit.. } } Kind regards } Valtcho Jeliazkov } } } } } }
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 13:37:09 2004
Dear Valtcho, I inherited the system in our University to manage the SEMs and TEM in our lab, but it has worked out well for me. 1. Because my department pays my expenses, users from my department do not pay for use. I have money available from a Shared Services account that all researchers in our department contribute to, to buy consumables like apertures and filaments. These users must still sign up on the calendar to reserve the instrument for themselves. 2. Other users from our university and other universities are charged a low hourly fee for use of all the instruments. I charge them $25 per hour for SEM and an additional $10 per hour if they use EDX. I also charge for gold coatings, film or other things that cost me money. 3. Commercial companies that want to use the instruments are charged commercial rates ( I charge $200 per hour). You need to find a commercial test firm that does SEM/EDX and see what their hourly charge is. Do not undercut them with a CFI grant-supplied instrument. I also set a priority system that spells out who has priority on the available time on the instrument. As an example: you would be first, researchers that supported your CFI application would be next, other researchers, Graduate Students, Undergraduate laboratories and projects would be defined. Commercial use is usually last, only if there is unused time available. All my instruments have a sign-up calendar so people can reserve them for a specified time. Some of my instruments are open time, so people can sign up for as much time as they want, when they want. The most popular (newest) instrument has a strict schedule where each day has four two-hour time slots and each user can only sign up for one slot. When they use that slot they can sign up for the next available slot. This gives everyone a chance. I hope this example gives you some ideas. Regards, Mary Mager Electron Microscopist Metals and Materials Engineering University of British Columbia 6350 Stores Road Vancouver, B.C. V6T 1Z4 CANADA tel: 604-822-5648 e-mail: mager-at-interchange.ubc.ca} -----Original Message----- } } From: Valtcho Jeliazkov (Zheljazkov) Ph.D. } [mailto:vjeliazkov-at-nsac.ns.ca] } Sent: Friday, March 19, 2004 11:52 AM } To: Microscopy-at-MSA.Microscopy.Com } Subject: [Microscopy] SEM/EDS system policy for usage } } } } } ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------ - } } Hi all, } } We have a new SEM/EDS system at NSAC. This is the } first electron microscope in our College. The } College administration has asked me to develop a } policy for usage, funding, prioritization and data } collection of the SEM/EDS unit. The } administration could not provide a technical } support for the unit. } I will be the major user of the system, I would } assume there will be between 10 and 15 other users } from the College. The system has been purchased } with CFI funding to support research programs. } } I would appreciate your comments and suggestions } on how to develop user fees, how much of my time } should be dedicated to the unit as teaching } others, managing the lab, and simple maintenance. } Perhaps I can ask the administration that 30-40% } of my time to be dedicated to the unit.. } } Kind regards } Valtcho Jeliazkov } } } } }
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 14:56:42 2004
I have some. I got it from out machine shop - when we had one. It is a (for SEM) large chunk and I can cut you a piece from it if nothing else easier turns up.
IMHO, The surface *must* be polished or the topo-contrast will swamp the Z-contrast. My GW Electronics, Type 47 system, "sees" the phases fine (No affiliation).
Regards, Woody
Woody White BWXT Services: http://www.bwxt.com/bwxt.html My Site: http://woody.white.home.att.net
-----Original Message----- } From: michael shaffer [mailto:michael-at-shaffer.net] Sent: Friday, March 19, 2004 10:47 AM To: Microscopy list; Sx50-Users
I'm evaluating backscatter detectors, and am in need of this familiar test material ... i.e., a difference in Z of 0.1 at Z=30. I see references to this standard in the supplies catalogs, but it is not immediately available, and the picture of it implies that it's not even polished ... which would contribute a lot to broadening the image's histogram peaks. It would help a lot if I could find a another source, and polish a sample of it next week. Is anyone aware of a poor mans' source of inclusion-free alpha-beta (or duplex) brass?
I don't even suppose it needs to be specifically AB brass, because all I need do is polish it extremely well, that it have neighboring phases to contrast, and have an average Z } 22 ... and make comparisons with the same material. For example, I have "pure" FeS exsolved in pyrrhotite, but the scale doesn't really allow me to select a relatively large area of each for determining the noise.
Suggestions most welcome ..
tia & cheerios ... shAf :o) Avalon Peninsula, Newfoundland www.micro-investigations.com
From MicroscopyL-request-at-ns.microscopy.com Fri Mar 19 16:26:53 2004
This is a second announcement for the workshop entitled “Multidimensional data presentation techniques” which will take place on April 4 from 12:30 to 4:30 just before the opening of the “Focus on Microscopy 2004” Conference in Philadelphia. Please hurry with your registration! There are only a few spots left.
For more information about the workshop please point you Internet browsers to:
http://www.cyto.purdue.edu/FOM2004/ - workshop web page or http://www.focusonmicroscopy.org/ - FOM2004 web page
This workshop will include an interactive tutorial on the use of a variety of techniques for multidimensional microscopy data presentation. Many advanced visualization packages for microscopists are commercially available. Similarly, plenty of applications for video processing and presentations can be found on the market. However, transforming complex data sets into the actual presentation for use in lectures or in web sites is not as easy as it seems. There are a variety of tricks-of-the-trade, useful suggestions, and some very nice inexpensive or free software obtainable. We would like to share our experiences and tell you about them!
You will learn how to present static 2D images as well as 3D datasets in the most efficient way. We will show you how to produce short animations using data from confocal/MP systems in highly compressed MPEG4-based formats. You will receive a handout and CD-ROM containing key materials presented in the workshop as well as a significant number of really valuable free utility software packages.
We will demonstrate: * How to compress microscopy data. What are the pros and cons of compression? Does it affect final results? What about lossy and lossless compression? * How you should present your 3D data. How to prepare 3D image reconstruction? How to create anaglyphs? How to protect the data in an anaglyph when you compress it? How to make a movie anaglyph/animation? * How to create an animation from a 3D construction. * How to create movies that are playable in PowerPoint, on web pages, or with other media. * How to understand codecs and their associated problems. * How to edit animations/movies using high-speed command line processing. * How to you add your name, logo of your institution, or other info into the movie. * How to deal with sound overlays. * How to reproduce a movie-making process. You will learn simple command line macros that are really fast!
If you are registered for the workshop, please take some time to take our pre-workshop survey. Your participation will help us greatly with the workshop preparation. We would like to know about your expectations, your level of experience in multimedia multidimensional data presentation techniques, and the issues you consider to be important. The link to the survey is present on the workshop web page.
Bartek Rajwa (rajwa at flowcyt.cyto.purdue.edu) Jennie Sturgis (jennie at flowcyt.cyto.purdue.edu) J. Paul Robinson (jpr at flowcyt.cyto.purdue.edu)
Purdue University Cytometry Laboratories Hansen Research Building 201 S. University Street West Lafayette, IN 47907 Telephone: (765) 494-0757 Fax: (765) 494-0517 Web: http://www.cyto.purdue.edu/
---------------------------------------------------------------------- The organizers of the workshop gratefully acknowledge the assistance of Media Cybernetics Corporation, the producer of Image-Pro Plus - an image analysis software package for fluorescence imaging, quality assurance, materials imaging, and various other scientific, medical, and industrial applications.
From MicroscopyL-request-at-ns.microscopy.com Sat Mar 20 08:48:56 2004
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (Elliotluke-at-yahoo.com) from http://www.msa.microscopy.com/MicroscopyListserver/MLFormMail.html on Friday, March 19, 2004 at 18:04:47 ---------------------------------------------------------------------------
Title-Subject: [Microscopy] [Filtered] Adventures in Amateur Micrography
Question: Dear Forum, I am a recent college graduate. I have a BS in Biology and hope to acquire my MS in microbiology in a few years. I enjoy microscopy and can not wait for graduate school to afford me the opportunities to stretch my microscopy muscles. I am looking to buy a microscope/microscopy setup for home use. I spend hours on the Internet looking at micrographs and associated photos. I would love to start taking my own photos and preparing slides in-house. I know that many seasoned microscopists share my passion for micrography. If you have any ideas on what sort of setup I should buy or ideas on how to approach amateur micrography on a budget please mail me. Thank you, Elliott
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (cynthia.green-at-njmoldinspection.com) from http://www.msa.microscopy.org/Ask-A-Microscopist.html on Saturday, March 20, 2004 at 06:44:14 ---------------------------------------------------------------------------
Email: cynthia.green-at-njmoldinspection.com Name: Cindy Green
Education: Graduate College
Location: City, State, Country
Question: I'm trying to locate a objective extender of about 10mm in length for a Wild DIN lens. Do you know of a good microscope parts source?
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (zachary_carr-at-eku.edu) from http://www.msa.microscopy.org/Ask-A-Microscopist.html on Sunday, March 21, 2004 at 14:41:54 ---------------------------------------------------------------------------
Email: zachary_carr-at-eku.edu Name: Zachary Carr
Organization: Eastern Kentucky University
Education: Undergraduate College
Location: Richmond, Ky, USA
Question: On a comparison microscope, how does the optical bridge split the image by means of mirrors?
Does anyone know how to accomplish a batch export of images from Autobeam in ISIS? I have 3000 odd images that need to be exported for processing on software that was written inhouse. Currently I have to open each image individually in Autobeam and then type/copy in a filename to export as tif.
Any help would be GREATLY appreciated.
Regards George
George Theodossiou Physicist / Electron Microscopist
AMCOR Research and Technology Ph: +61 3 9490 6135 Fax: +61 3 9499 4295 Mobile: 0409 568 840 email: George.Theodossiou-at-amcor.com.au
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From MicroscopyL-request-at-ns.microscopy.com Mon Mar 22 07:27:05 2004
As with any material there are several ways of specimen preparation for observation of tooth structure. Specimens could be fractured or polished, they could be etched with acids. For observation of an enamel hi vac mode is usually better.
It is difficult to advise you without knowing the specifics of the research. You are welcome to contact me off line for more discussion.
Vladimir
Vladimir M. Dusevich, Ph.D. Electron Microscope Lab Manager 3127 School of Dentistry 650 E. 25th Street Kansas City, MO 64108-2784
} Email: hadden-at-wingate.edu } Name: Lee Hadden } } Organization: Wingate University } } Title-Subject: [Microscopy] [Filtered] MListserver: } } Question: One of my SEM students is working with teeth. Can } anyone recommend the best way to view the teeth in the SEM, } i.e. hi vac SEI or low vac BEI [or any other suggestions for } most effective imaging of enamel structure]? We are using a } JEOL LV5600 SEM. } } Thanks in advance. } } Lee Hadden } Department of Biology } Wingate University } Wingate, NC 28174 } } -------------------------------------------------------------- } ------------- } }
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 22 12:03:00 2004
Zachary, Here is a very quick explanation of the light path of the comparison microscope. As light comes up through the objective lens above each of the two stages it enters a set of prisms that erect the image and redirect it toward the center of the bridge. You could think of this set of prisms as a mirror set at 45 degrees to reflect the image at a right angle toward the center of the bridge, the prisms do more, but this will give you an idea of what is happening. At the center of the bridge is another set of prisms that reflect the image up to the eye pieces. You could think of this prism set as two mirrors set back to back at 45 degrees each, with the tops of the mirrors meeting at a very sharp edge, they redirect both fields (one from each objective) 90 degrees upward to the eye pieces. In some scopes this center prism set is on a moveable mount that lets you move this set of prisms into position to allow 50% of the field from each objective, all of one field and none of the second, or any percent of each that makes up 100%, for example 70% of one field and 30% of the other. Here is a very rough diagram of the light path. If you want a better one I think I have a photo of a part of a Leica poster with a diagram of the UFM 4's optical path, I could send it off the list but don't want to clutter the list with attachments. Drop me a note if you need this, I'll see if I can find it or snap another.
| /------/\------\ | | _ _
Jim
James L. Roberts Forensic Scientist / Firearm and Toolmark Examiner Ventura County Sheriff's Forensic Science Laboratory 800 S. Victoria Ave. Ventura, CA. 93009
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (zachary_carr-at-eku.edu) from http://www.msa.microscopy.org/Ask-A-Microscopist.html on Sunday, March 21, 2004 at 14:41:54 ---------------------------------------------------------------------------
Email: zachary_carr-at-eku.edu Name: Zachary Carr
Organization: Eastern Kentucky University
Education: Undergraduate College
Location: Richmond, Ky, USA
Question: On a comparison microscope, how does the optical bridge split the image by means of mirrors?
M&M 2004 Attendees there will be two additional short courses offered at M&M2004 this year that were not listed in the call for papers. Their descriptions are listed below. You can sign up for these courses at the meeting or online.
Short Course X18
Title: Microspectroscopy - Raman, FT-IR and EDXRF Microscopy
9:00am to 5:00pm
Instructors: Fran Adar, John Reffner, Paul Dinh
Today more and more laboratories are combining microscopy with the power of spectroscopy to provide a detailed characterization of their samples. This short course will show how laboratories are using Raman, FT-IR and EDXRF Microspectroscopy to their benefit. These three techniques provide molecular and/or elemental information with a spatial resolution at the micron level. They can also be used for hyperspectral imaging to show how the molecular and elemental structure changes throughout a sample. Attendees will leave the course with a knowledge of the varied uses of the instrumentation and the applicability of these methods to their particular materials. Examples from biological, pharmaceutical, polymer, semiconductor and forensic applications will be used to illustrate the power of these techniques. The course will offer lectures, hands on instrumentation and a chance for questions. Participants are welcome to bring samples.
Short Course X19
Title: Application Pathways- Native sample to Immunolabeling via Tokayasu and High Pressure Freezing
9:00am to 5:00pm
Instructors: Kent McDonald & Jan Slot
High pressure freezing is the way to localize or characterize organelles, subcellular components and gene products in electron microscopy. This single-day short course will discuss the process of sample preparation using cryo techniques.
The course will review methods for cryopreparation of biological samples. High pressure freezing, why,? the advantages of HPF over chemical and or microwave fixation. After HPF what techniques or methods of tissue examination can be used, such as cryo planing, cryosectioning, freeze substitution, freeze fracture and immunolabeling. The course will offer lectures, hands on of related instrumentation and roundtable discussions.
-- John Mansfield PhD MInstP North Campus Electron Microbeam Analysis Laboratory 417 SRB, University of Michigan 2455 Hayward, Ann Arbor MI 48109-2143 USA Phone: (734) 936-3352 FAX (734) 763-2282 Cell. Phone: (734) 834-3913 (Leaving a phone message at 936-3352 is preferable to 834-3913) Email: jfmjfm-at-engin.umich.edu URL: http://emalwww.engin.umich.edu/people/jfmjfm/jfmjfm.html Location: Lat. 42° 16' 48" Long. 83° 43' 48" AIM: thejfmjfm
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 22 13:32:56 2004
We (an hourly under my direction) wrote a utility to export batches of ISIS Autobeam images to TIFF format. We have used it on hundreds of images. I don't think we are up to the thousands yet. You can download the package from ftp://www.marl.iastate.edu/Utilities/ISIS-TIFF/. Documentation is included with the executable files.
I think we had some problems where the last image of the selected set might not get converted. If so, you may need to convert it after the larger batch. But I think it has been working okay of late, especially when I convert the entire job.
As long as you remember you get what you pay for, you may drop me a line if you have detailed questions about its use.
At 01:44 AM 3/22/2004, George Theodossiou wrote:
} Hi, } } Does anyone know how to accomplish a batch export of images from Autobeam in } ISIS? I have 3000 odd images that need to be exported for processing on } software that was written inhouse. Currently I have to open each image } individually in Autobeam and then type/copy in a filename to export as tif. } } } Any help would be GREATLY appreciated. } } Regards } George } } George Theodossiou } Physicist / Electron Microscopist } } AMCOR Research and Technology } Ph: +61 3 9490 6135 } Fax: +61 3 9499 4295 } Mobile: 0409 568 840 } email: George.Theodossiou-at-amcor.com.au
------------------------------------------- No files should be attached to this message ------------------------------------------- Warren E. Straszheim, Ph.D. Materials Analysis and Research Lab Iowa State University 46 Town Engineering Ames IA, 50011-3232
I have used an AGFA Duoscan 1200 for some years for TEM negatives. Recently it stopped working and it seems that it is easier to buy a new one than repair the old machine.
Can anyone recomend a suitable (inexpensive) machine? As far as my experience goes, the low end scanners do not give acceptable results with dense negatives. The Duoscan was acceptable perhaps due to better sensitivity. More sensitive scanners may become too expensive for our lab. So, I'm looking for a suitable compromise: enough sensitivity v. lowest price
Thanks in advance
A.P. Alves de Matos Biologist Electron Microscopy Unit Curry Cabral Hospital, Lisbon
From MicroscopyL-request-at-ns.microscopy.com Mon Mar 22 22:20:46 2004
This is the second announcement for the upcoming free, 2-day "miniworkshop" in Cryoultramicrotomy for Materials Sciences that will be held at West Chester University of Pennsylvania.
If you have not yet sent your RSVP, please do so to any of the people listed below. We need an accurate count of attendees for meals and refreshments.
The details are as follows:
Members who have an interest in materials science are cordially invited to a free two-day "mini-workshop" on cryoultramicrotomy for the materials sciences. This topic is of special interest for those who work with polymers or other materials which could benefit from ultrathin sectioning or surface "polishing" at low temperatures (no embedding required).
This invitation is extended to persons who are located in the greater Philadelphia/Baltimore/Washington, D.C./New York City area. The workshop will be hosted by the Center for Advanced Scientific Imaging (CASI) at West Chester University of Pennsylvania.
**What**: Mini-Workshop on Cryoultramicrotomy for Materials Science
**When**: Tuesday, March 30, 2004, 11:00 am through Wednesday, March 31, 2004, 4:00 pm.
Visitors are invited to arrive early on Tuesday, March 30 for tours of the Center for Advanced Scientific Imaging. The lab will be open at 8:00 am on Tuesday.
**Where**: West Chester University of Pennsylvania, Schmucker Science Center South, Room SSS017. Schmucker Science Center South is located on the corner of South Church Street and Rosedale Avenue in West Chester, PA. West Chester is located approximately 30 miles from Philadelphia.
**Format**: A presentation on cryoultramicrotomy and its applications in the materials sciences will be given on the first day, as well as demonstrations on the preparation of cryotools (hair probes, large and small wire loops, etc.) and glass knife making and evaluation. The care and cleaning of diamond knives will also be discussed.
The demonstrations will be followed by open lab sessions for the attendees to prepare their own cryotools and glass knives.
Also on the first day an introduction to the cryoultramicrotome will be given, and attendees will have an opportunity for hands-on use of the instrument.
The second day will be reserved for attendees to sign up in small groups for additional time and training on the instrumentation, depending on the attendees' individual needs.
**Limited Space Available**: The lectures and demonstrations on the first day are open to everyone, but space is limited to *10 persons* for the in-depth training and extended use of the instrumentation on the second day.
**Contacts**: To RSVP or to reserve a seat for the second day's sessions, please contact any of the following people:
Dr. Fred Monson, CASI / West Chester University of Pennsylvania, 610.738.0437, {fmonson-at-wcupa.edu}
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (jrobson-at-rdg.boehringer-ingelheim.com) from http://www.msa.microscopy.org/Ask-A-Microscopist.html on Tuesday, March 23, 2004 at 06:41:11 ---------------------------------------------------------------------------
Email: jrobson-at-rdg.boehringer-ingelheim.com Name: John Robson
Organization: Boehringer Ingelheim
Education: Undergraduate College
Location: Danbury, CT
Question: I've been asked to perform several measurements on a polyethylene part. I need to determine the length and diameter (at several points) of a channel that is roughly 0.3mm wide X ~2.0mm long. Years ago we had worked with latex casting materials for this purpose unfortunately shrinkage was a constant issue. I would appreciate any recommendations on alternative casting materials that would allow us to accurately reproduce an impression of the part with minimal shrinkage. Thank You, John.
O.k., folks the radiation safety people here are looking at re-classifying *ALL* the uranyl compounds on campus as lisenced materials (i.e. radioisotopes) with all the requirments and certifications that go along. As careful dropwise users of Uranyl acetate I am trying to avoid unneeded headaches, and trying to get low concentration low quantity Uac to be considered separately from the high quantity chemistry users. To that end I am collecting info on how other institutions handle it.
So folks how does your institution classify Uac: Radioisotope or not. And how is it disposed of?
Especially Ohio folks appreciated! Thank you.
(Note: I have searched the listserv archive 2001-2004 before posting this, and we've only discussed it a little in early Jan 2004)
==============
Since some folks may ask: How we currently use and handle UA:
The EM Facility does use Uranyl Acetate on a regular basis for heavy metal "staining" of transmission electron microscope (TEM) samples. We work with 0.5% & 2.0% aqueous solutions (approximately 80-85% of the work uses 0.5% UAc), and we use ~ 0.3-0.6g total per year. The EM grade Uranyl acetate for staining is sold as a "depleated" Uranyl acetate (since we need the heavey metal and not the radiation), and is difficult to detect above back ground radiation. We use 10ul drops at a time, these droplets are collected and stored in lable waste containers, and rinse water is also collected in the waste containers (Containers only contain Uac and water). Transfer pipettes are rinsed before disposal. Waste containers are collected by EHS regularly. We treat the Uac as a serious heavy metal toxin as we do our other heavy metal stains (Pb, Os, W, Cr, Ag, etc.).
If the decision is made to consider *ALL* Uranyl compunds licensed material, the this will have a serious impact on the facility to meet the licensing requirments, for very little increased safety. One of our standing polcies at present is *NOT* to allow radioisotopes in the facility as we are not certified to handling them. We would need radiation training of all TEM users. We would need locked storage of stocks, working solutions, and wastes (since locking the facility would not be practical), then users would be facing moving solutions from the locked location to the application site, and back again. Whereas at present the working solution and wasters containers used are not moved they are simply opened and closed. In this case I think licensing would increase the risk (due to dropping and spillage) rather than decrease.
I would just ask that the radiation safety committee perhaps consider subcategories of Uranyl compounds and/or quanities used.
=====================
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 350 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu http://www.emf.muohio.edu
"RAM disk is NOT an installation procedure."
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 23 08:03:57 2004
I too, had a limited budget and a need for a scanner. A few years ago, I bought an Epson Expression 1600 Professional scanner with trans-illumination. I've been pretty happy with it. It comes with holders for negatives, but for the 3.25 x 4 inch negs I have. I place my negatives directly on the glass and use the focus setting for contact and I haven't had problems. (OK everyone, a collective gasp of horror here...but really, its been OK). Lee -- Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Optical Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 23 08:30:49 2004
We bought the Epson Perfection 3200 Photo and it works well. We also have to put our TEM negs directly on the glass, but we have had no problem with it.
Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 55 Fruit Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) msherwood-at-partners.org
-----Original Message----- } From: AP Alves de Matos [mailto:apamatos-at-oninet.pt] Sent: Monday, March 22, 2004 5:34 PM To: Microscopy-at-MSA.Microscopy.Com
Dear Colleagues
I have used an AGFA Duoscan 1200 for some years for TEM negatives. Recently it stopped working and it seems that it is easier to buy a new one than repair the old machine.
Can anyone recomend a suitable (inexpensive) machine? As far as my experience goes, the low end scanners do not give acceptable results with dense negatives. The Duoscan was acceptable perhaps due to better sensitivity. More sensitive scanners may become too expensive for our lab. So, I'm looking for a suitable compromise: enough sensitivity v. lowest price
Thanks in advance
A.P. Alves de Matos Biologist Electron Microscopy Unit Curry Cabral Hospital, Lisbon
From MicroscopyL-request-at-ns.microscopy.com Tue Mar 23 08:52:20 2004
We also use the EPSON 3200 Photo, with great results. If better scans (for enlargements) need to be performed, we use the NIKON SuperCoolScan 8000 ED which gives 4000 ppi images to work with. The slowness is the reason for only using the NIKON for specific negatives, BUT as for the resolution, and the contrast depth - it is unsurpassed. Stan
"Sherwood, Margaret" wrote:
} ------------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ------------------------------------------------------------------------------- } } We bought the Epson Perfection 3200 Photo and it works well. We also have } to put our TEM negs directly on the glass, but we have had no problem with } it. } } Peggy Sherwood } Lab Associate, Photopathology } Wellman Laboratories of Photomedicine (W224) } Massachusetts General Hospital } 55 Fruit Street } Boston, MA 02114 } 617-724-4839 (voice mail) } 617-726-6983 (lab) } 617-726-3192 (fax) } msherwood-at-partners.org } } -----Original Message----- } } From: AP Alves de Matos [mailto:apamatos-at-oninet.pt] } Sent: Monday, March 22, 2004 5:34 PM } To: Microscopy-at-MSA.Microscopy.Com } Subject: [Microscopy] TEM Acquiring new scanner for negatives } } ---------------------------------------------------------------------------- } -- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- } http://www.msa.microscopy.com/MicroscopyListserver } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------------- } --- } } Dear Coll