To All, } } The company I work for currently utilizes a coated glass slide that is } amine reactive for printing our microarrays. The supplier of the } substrate is reticent to divulge any information regarding the chemical } composition, thickness, and QC measures employed in their process. } Subsequently, we would like to ascertain as much info as possible } regarding the coating thickness, surface topography, etc. and were } hoping that experts in the field of microscopy might offer some } suggestions or be willing to undertake such a project. } } } Chris Michaelson }
Hi- we are in the process of purchasing a new confocal, and are looking at a Zeiss510 Meta or a Leica AOBS or SP2. I think that in the demo process we are seeing what we need to, and are impressed by the capabilities of each. I would appreciate user perspectives that we cannot get from the manufacturers though- are there instrument aspects of either microscope that are repeatedly problematic or limiting? Do multi-user facilities have particular difficulty with either? I do not expect an obvious "winner" for an answer, but am more interested in hearing what some of the ups and down are, and seeing if those problems or circumstances would be issues for us. Thanks for any experience you an offer, I am sure it will be useful.
Betsey ************************************************************************ Betsey Pitts Research Associate/Facilities Manager, Microscopy The Center for Biofilm Engineering
366 EPS Building, Montana State University Bozeman, MT 59717
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I always understood this to be due to "photons is photons". Granted that the energy of an IR photon is much less than an x-ray photon. (What is the wavelength cutoff for generating an electron-hole pair?) However, there are just so darn many of those IR photons. So not only do we get the very low energy counts due to the photons, but also we get a lot of dead-time and pile-up of photons. I usually see our dead time max out when the light comes on. It takes several seconds for the circuitry to get back to normal after we shut the light off.
I know x-ray windows are supposed to be treated to render them opaque to infra-red. However, that treatment appears to be more effective in some cases than other. Both of our detectors are affected at least some when the cameras are turned on.
Regarding cathodoluminescence, I wonder if the signal is too weak to be of concern. A 20-kV beam at 1 nA is only dumping 20 micro-watts of power into the sample. Only a small fraction of that gets converted over to light. I don't know the power rating of our IR bulb on our chamber scope, but I am sure it is quite a few orders of magnitude more powerful.
Warren
At 10:51 PM 1/30/03 +0000, you wrote:
} Don't know, but I would be interested to know. We need a physicist } here! } I am also intrigued to note that some of my specimens show } cathodoluminescence outputs sufficiently bright for the position of } the scanning beam to } be visible on the chamber cam. } How bright does it need to get to have an impact on counts, and what } artefacts might result? } } Chris } } ----- Original Message ----- } From: "Ritchie Sims" {r.sims-at-auckland.ac.nz} } To: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} ; } {microscopy-at-sparc5.microscopy.com} } Sent: Thursday, January 30, 2003 9:38 PM } Subject: IR LEDS/EDS } } } } I had a problem like this. } } } Turned out to be caused by the infra-red leds of my chamber } camera. } } } When the camera is switched off, the counts (3-5k per second) } return } } } close to zero. } } } } } } Chris } } } } } } } What would be the mechanism of this effect, I wonder? } } } } rtch } } } } Ritchie Sims Phone : 64 9 3737599 ext 7713 } } Department of Geology Fax : 64 9 3737435 } } The University of Auckland email : r.sims-at-auckland.ac.nz } } Private Bag 92019 } } Auckland } } New Zealand
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More than 25 years ago we had our machine shop make a rectangular box and cover of sheet stainless steel, complete with handles. It holds both a loaded film box and an exposed film box from a JEOL 100 CX. It still looks and works like new and was worth the expense.
Bernie Kestel Materials Science Division Argonne National Laboratory 9700 South Cass Avenue Argonne, Il., 60439
by sparc5.microscopy.com (8.9.3+Sun/8.9.3) id AAA13285 for dist-Microscopy; Sun, 2 Feb 2003 00:37:11 -0600 (CST) Received: from njz_spm_filter (sparc5 [206.69.208.10]) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) with SMTP id AAA13207 for "MicroscopyFilteredEmail3-at-msa.microscopy.com"; Sun, 2 Feb 2003 00:36:40 -0600 (CST) Received: from mailgw.solutia.com (mailgw.solutia.com [205.191.166.80]) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) with ESMTP id AAA13164 for {microscopy-at-sparc5.microscopy.com} ; Sun, 2 Feb 2003 00:36:24 -0600 (CST) Received: from mail pickup service by mailgw.solutia.com with Microsoft SMTPSVC; Sun, 2 Feb 2003 01:30:59 -0500 Received: from soidmzvw01.solutia.com ([205.191.64.37]) by mailgw.solutia.com with Microsoft SMTPSVC(5.0.2195.5329); Sat, 1 Feb 2003 01:02:36 -0500 Received: from 206.69.208.10 by soidmzvw01.solutia.com (InterScan E-Mail VirusWall NT); Sat, 01 Feb 2003 00:58:00 -0500 Received: (from daemon-at-localhost) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) id OAA02019 for dist-Microscopy; Fri, 31 Jan 2003 14:11:01 -0600 (CST) Received: from njz_spm_filter (sparc5 [206.69.208.10]) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) with SMTP id OAA01995 for "MicroscopyFilteredEmail3-at-msa.microscopy.com"; Fri, 31 Jan 2003 14:10:30 -0600 (CST) Received: from dragon.ti.com (news.ti.com [192.94.94.33]) by sparc5.microscopy.com (8.9.3+Sun/8.9.3) with ESMTP id OAA01982 for {microscopy-at-sparc5.microscopy.com} ; Fri, 31 Jan 2003 14:10:05 -0600 (CST) Received: from dlep11.itg.ti.com ([157.170.133.18]) by dragon.ti.com (8.12.6/8.12.6) with ESMTP id h0VK4cfE009221; Fri, 31 Jan 2003 14:04:38 -0600 (CST) Received: from dlep11.itg.ti.com (localhost [127.0.0.1]) by dlep11.itg.ti.com (8.9.3/8.9.3) with ESMTP id OAA15460; Fri, 31 Jan 2003 14:04:37 -0600 (CST) Received: from ti.com (asp0142618.sc.ti.com [156.117.194.48]) by dlep11.itg.ti.com (8.9.3/8.9.3) with ESMTP id OAA15456; Fri, 31 Jan 2003 14:04:37 -0600 (CST) Message-ID: {3E3AD7AC.CC885713-at-ti.com}
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Chris: light shining on a semiconductor (which the detector is) generates electron-hole pairs, just like an x-ray does. But the magnitude of the number of these pairs is much higher because there are usually more photons in a light beam than characteristic x-rays being generated by a sample. This flood of electron-hole pairs overwhelms the amplifier and the counting electronics. Some detectors have metallized (mere atoms of metal, so they won't interfere w/ analysis) windows to keep out the 'ambient' light, such as maybe generated by CL. But if your chamberscope's light source is oriented so that it shines directly on the detector, enough light can get in to cause problems. The collimator on the detector nose can also help keep light out. I've worked on SEMs that have had the chamberscope light source directly opposite the detector and had to be vigilant about turning it off when is was not needed. I now have a scope that has the light source behind the detector and it give no trouble at all. For the curious, on the Oxford Instruments site, on the page for their EDX hardware (http://www.oxford-instruments.com/ANLPDP174.htm) near the bottom right hand side is a list of "Related PDFs" and the last one is "EDX Hardware Explained". If you've ever wondered what's in the snout of a detector and how it works its magic, this is a good read. NOTE: I have no interest, financial or otherwise, in Oxford Instruments. Just a satisfied customer.
Chris Jeffree wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Don't know, but I would be interested to know. We need a physicist } here! } I am also intrigued to note that some of my specimens show } cathodoluminescence outputs sufficiently bright for the position of } the scanning beam to } be visible on the chamber cam. } How bright does it need to get to have an impact on counts, and what } artefacts might result? } } Chris } } ----- Original Message ----- } From: "Ritchie Sims" {r.sims-at-auckland.ac.nz} } To: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} ; } {microscopy-at-sparc5.microscopy.com} } Sent: Thursday, January 30, 2003 9:38 PM } Subject: IR LEDS/EDS } } } } } } } } } I had a problem like this. } } } Turned out to be caused by the infra-red leds of my chamber } camera. } } } When the camera is switched off, the counts (3-5k per second) } return } } } close to zero. } } } } } } Chris } } } } } } } What would be the mechanism of this effect, I wonder? } } } } rtch } } } } Ritchie Sims Phone : 64 9 3737599 ext 7713 } } Department of Geology Fax : 64 9 3737435 } } The University of Auckland email : r.sims-at-auckland.ac.nz } } Private Bag 92019 } } Auckland } } New Zealand } }
-- ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Becky Holdford (r-holdford-at-ti.com) 972-995-2360 972-648-8743 (pager) SC Packaging FA Development Texas Instruments, Inc. Dallas, TX ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
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Peter, In that case, a TEM would also be an electron microprobe. I think it is better if the names are used specifically. Electron microprobes are designed quite differently from SEMs. Call an SEM with WDS an SEM with WDS.
At 11:12 AM 1/30/2003 -0500, Peter Tomic wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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Dear Ritchie, I believe the Si(Li) detector is sensitive to visible light, as it is to the x-ray photons. With a Be window the visible light could not penetrate to the detector, but with the new thin membrane windows the IR gets right through. 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: "Ritchie Sims" {r.sims-at-auckland.ac.nz} To: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} ; {microscopy-at-sparc5.microscopy.com} Sent: Thursday, January 30, 2003 1:38 PM
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Gary I think, you wrong: fragmented file occupied more physical space on HD (yes) but fragmented/non-fragmented files has the same bit-size. So fragmentation may affect the reading/writing time only, which on the modern computers is negligibly. If the disk seriously fragmented, yes, it may affect computer's performance in general. "Save as" command not necessary writes in non-fragmented space. NTFS usually decently manage this issue and keep fragmentation at relatively low level, but it's OS decision where to place your file (for instance, if file is small, it would be placed in the NTFS analog of FAT at the beginning of drive- this is special precaution against fragmentation). If the HD is seriously fragmented, even "Save as" will cause the file fragmentation. Sergey
At 08:18 AM 1/31/2003, 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 Box 951737 Los Angeles, CA 90095-1737
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We use tin cookie boxes. They come in all different sizes and before christmas, 2001 we asked regular lab users to save their cookie boxes if they received any as christmas gift. They are light tight and so far have been working great.
By the way, we just asked for used, empty boxes, no cookies.
Soumitra
************************************************************* Soumitra Ghoshroy College Associate Professor, Biology Director, Electron Microscopy Lab Box 3EML New Mexico State University Las Cruces, NM 88003 Tel: 505-646-3268 (office), 646-3283 (lab) Fax: 505-646-3282 e-mail:sghoshro-at-nmsu.edu URL:http://confocal.nmsu.edu/eml
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I think you would have to worry about two problems - (1) shrinkage during the actual polymerization step and (2) compression during thin sectioning. I don't have the reference off the top of my head but Daniel Studer has a nice paper (J. Microscopy about 1-2 years ago, I think) in which he shows significant (up to 50% if i remember correctly) along the cutting axis. his paper showed this could be avoided using an oscillating diamond knife. this paper has important implications for high resolution measurements since the compression would be different in the different axes.
At 10:08 AM 1/31/2003 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Associate Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
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Dear Jim, I use the black plastic bags that the photographic paper comes wrapped in to transport my film boxes to the dark room. I discovered the hard way that the boxes themselves were not light tight. 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: "Jim Romanow" {bsgphy3-at-uconnvm.uconn.edu} To: {microscopy-at-sparc5.microscopy.com} Sent: Thursday, January 30, 2003 1:36 PM
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On Friday, January 31, 2003, at 01:54 AM, Ji, Ying wrote:
} I am doing a double tilting experiment with TEM. From diffraction zone } A to } diffraction zone B, I tilted X axis for xx degree and Y axis for yy } degree. } Could anyone let me know how to calculate the angle between diffraction } zones A and B. } } Thank you very much in advance! } Dear Yun, I don't have my spherical trigonometry book here, having moved recently, but the way to approach the problem is to imagine the electron beam incident on the North pole with the Greenwich meridian facing you, then perform the tilts and locate the position of the incident beam after these have been done. If you have a double tilt holder like the one I used when I was in Albany NY, the tilts can be done in either order, so tilt in Y (keeping the Greenwich meridian facing you) so the beam is now incident on latitude 90-yy, then tilt through xx degrees along the great circle perpendicular to the Greenwich meridian and passing through it at latitude 90-yy. You will have a right spherical triangle; the hypotenuse is the angle you're looking for. Yours, Bill Tivol 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
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As I understood it at M&M 2002 in Quebec City, the Diatome oscillating knife was close to market, as in the 'fine-tuning' stage. You might want to inquire of Diatome US as to the state of that knife. The reduction in compression for soft materials was phenomenal.
Tom Malis
Dr. Tom Malis Scientist Advisor Natural Resources Canada Govt. of Canada 613-995-7358 malis-at-nrcan.gc.ca
-----Original Message----- } From: Tom Phillips To: Nahirney, Patrick (NIH/NIAMS) Cc: Microscopy-at-sparc5.microscopy.com Sent: 1/31/2003 11:49 AM
I think you would have to worry about two problems - (1) shrinkage during the actual polymerization step and (2) compression during thin sectioning. I don't have the reference off the top of my head but Daniel Studer has a nice paper (J. Microscopy about 1-2 years ago, I think) in which he shows significant (up to 50% if i remember correctly) along the cutting axis. his paper showed this could be avoided using an oscillating diamond knife. this paper has important implications for high resolution measurements since the compression would be different in the different axes.
At 10:08 AM 1/31/2003 -0500, you wrote: } ----------------------------------------------------------------------- - } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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Light tight container for film transport?
Black plastic bags win the popularity (and economy) contest!
Quick list of light tight container suggestions:
Black plastic photo paper bag with or without cardboard box Ammunition storage box Paint can Pelican camera equipment case Spring loaded paper safe box Plastic tool/tote box Custom made stainless steel box
Thank you very much for all of the feedback. I am leaning toward the ammo box because it might hold up well under student abuse; the most bullet proof solution:}
Regards, Jim
James S. Romanow The University of Connecticut Physiology and Neurobiology Department Electron Microscopy Facility Unit 2242 354 Mansfield Road Beach Hall, Room 129 Storrs, CT 06269-2242
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Dear Fellow Microscopists,
A colleague of mine recently asked about a type of "Laser Scanning Phase Contrast Microscopy", for which I do not have much knowledge.
Please advise: What's the major difference between this and the conventional LSCM that we use? Which institution in the East Coast might have such a facility?
Any advice is highly appreciated and will be forwarded to this colleague. Have a great weekend!
QC
Qian-Chun Yu, MB, Ph.D. Director Biomedical Imaging Core Laboratory University of Pennsylvania Department of Pathology & Lab Medicine School of Medicine 110 Richards Building Philadelphia, PA 19104
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Please help. I'll pay any price for a decent copy of a Reichert OM-U3 ops/service manual. Someone was nice enough to send a copy of a copy, of a copy, etc., but all the illustrations are illegible. That was an ops manual, and I'll still need a service manual, or at least a schematic or wiring diagram. Anyone with some spare specimen block holders would be helpful also. The instrument that I have only came with one flat block holder. I have lots of lab stuff I would be happy to trade.
--
My address is: Equipment Resurrection 1005 Terra Nova Boulevard, Suite 2 Pacifica, CA 94044. The phone number is 650-738-0351 and web address, http://equiprx.net/.
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Chris Jeffree wrote:
} } Don't know, but I would be interested to know. We need a physicist } here! } ----- Original Message ----- } From: "Ritchie Sims" {r.sims-at-auckland.ac.nz} } To: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} ; } } } } } What would be the mechanism of this effect, I wonder? } } } } rtch }
I expected the usual suspects to leap on this, but since it hasn't happened, I will chime in. I are an engineer, not a physicist; take with the appropriate grain of salt, but I have worked in pulse processor design.
The Al coatings on thin window detectors do a good job of keeping out visible light, but are fairly transparent to IR. The IR photons are absorbed just like X-rays, but because their energy is so low and the flux high compared to X-ray photons, the result is a nearly continuous flow of current through the detector which is indistinguishable (to the pulse processing electronics) from a "leaky detector". Essentially, high leakage current raises the noise threshold setting required to prevent false triggering of the pulse processor. Since thin-window detectors are intended to trigger on low-energy X-rays, that threshold in a properly funtioning detector is set fairly low. Therefore, the IR- induced "leakage current" causes the pulse processing electronics to start triggering continuously on noise, causing a large peak at the low end of the spectrum.
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We're having an odd problem.
We're staining COS cells with Alexa 647 phalloidin (Mol. Probes A-22282 lot 41B1-1). The f-actin is staining fine and looks great when we excite at 633 nm and detect through the Leica AOBS or normal Cy5 filter set.
However, when we excite at 488 nm, we're seeing the f-actin staining appearing in the green channel too. We checked on different microscopes.
Has anybody else seen something as weird as this? We suspect contamination, but don't know where it would have come from unless it is an isomer or something from the manufacturing process.
One of our controls is the Alexa-phalloidin staining alone without any antibodies, so we know it's not some weird antibody binding artifact.
Any help appreciated.
Thanks.
____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
the resultant y" axis orientation (equivalent to the specimen normal in this case) is given as:
y" = -x sin(a)cos(b) + y cos(a)cos(b) + z sin(b)
The total tilt is found from the arccos of the dot product of the initial y axis and the final y" axis, which gives (assuming unit vectors)
cos(theta) = cos(a)cos(b)
which is what I stated this morning.
Dick Fonda
At 9:54 AM +0000 1/31/03, Ji, Ying wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Those of you in the path of the debris feild of the tragic break up of the space shuttle Colombia have a unique opportunity to try to catch microscopic debris from it over the next few days as the winds move it over the south east part of the United States. This is the largest event of its kind and first chance for this kind of research.
Pans of glycerin, water or some other liquid or possibly the sticky side of tap or some kind of gel to trap the particles on roofs and in the open at ground level should catch this debris. Some kind of baffles to protect the pans from wind should help catch small particles and protect from contamination from the surrounding area.
Reports in California by an astronomer of small flashes following the shuttle as it pass over may extend the area were debris can be found.
The current winds aloft should carry the debris along the Gulf Coast and across central Florida if they continue as they are now.
Gordon Couger gcouger-at-couger.com
I collect links on information related to light microscopes. http://www.couger.com/microscope/links/gclinks.html Please forward any links or information you think might be useful to others.
I'm trying to set up the column alignment of my JSM 840, and rotation of the Y stigmator gives such a lot of image shift that it's hard to use. The X stig gives almost none.
There are trim pots which balance the currents to the stig coils, but the amount of image shift is way more than can be corrected for by the trim pots.
The currents flowing in all of the 8 coils (4 'X', 4 'Y') are all very similar, and they all change similarly when the stig controls and the trim pots are turned, so it seems that the stig electronics and the coils are all OK.
Is this effect likely to be caused by a gross misalignment of the column?
Any tips on how to remedy?
thanks
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
In my last order of Kodak 4489 film, the boxes were marked "new formulation". Since it has been some time since I've needed to order film, I was wondering if any of you have noticed sufficient differences in exposure, density, etc. with this new film that required re-calibration of the photographic parameters on your microscopes. The last time Kodak re-formulated their emulsion, we needed to do extensive testing and calibration of our TEMs to obtain photos with the desired exposure levels.
Thanks.
Valerie M. Knowlton Research Assistant/Teaching Technician Center for Electron Microscopy 1219 Gardner Hall, Box 7615 North Carolina State University Raleigh, NC 27695
Richard Fonda is right. I made a foolish mistake when multiplying the two rotation matrixes together.
My sincerest apologies. Paul
======================= Paul Baggethun Engineer Alcoa Technical Center Alcoa Center, PA 15069 USA =======================
-----Original Message----- } From: Richard W. Fonda [mailto:fonda-at-anvil.nrl.navy.mil] Sent: Friday, January 31, 2003 4:20 PM To: Ji, Ying; 'microscopy-at-sparc5.microscopy.com'
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There was some debate about the answer I gave to this question this morning, so I looked into the math a bit further.
If you consider the standard rotation of cartesian coordinates about the z axis by an angle, a, as follows:
x' = x cos(a) + y sin(a) y' = -x sin(a) + y cos(a) z' = z
and follow this by a rotation about the x' axis by an angle, b:
the resultant y" axis orientation (equivalent to the specimen normal in this case) is given as:
y" = -x sin(a)cos(b) + y cos(a)cos(b) + z sin(b)
The total tilt is found from the arccos of the dot product of the initial y axis and the final y" axis, which gives (assuming unit vectors)
cos(theta) = cos(a)cos(b)
which is what I stated this morning.
Dick Fonda
At 9:54 AM +0000 1/31/03, Ji, Ying wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Greetings Betsey, It is a wise idea to contact confocal facilities and get some input regarding experiences with established instrumentation. Another forum you may wish to query is the Confocal Listserver {confocal-at-listserv.buffalo.edu} , and you may wish to look at the archives (http://listserv.acsu.buffalo.edu/cgi-bin/wa?S1=confocal); there was a pertinent thread discussing the two platforms last week. There is a listserver set up as a resource for administrators of Leica CFM equipment as well which you may wish to address--subscribers include 60 or so facility coordinators around the globe who manage Leica confocal microscopes. If you would like I can forward your query to this listserver (you have to be subscribed to post as an anti-spam measure). Lastly, you might try to find facilities on the web (eg. enter "Leica SP-2" into a Google search and see what comes up). Most facilities are happy to share their experiences. No high tolerance instrument will perform without incident indefinitely and intelligent purchase decisions take this into account. Sophisticated platforms such as the SP-2 and the META showcase synergy between a number of instrument sub-systems; seemingly minor problems or mis-calibrations in the imaging pipeline can wreak havoc with important measurements. It is important that problems can be addressed effectively and efficiently--for this it is necessary to have sincere support from a particular manufacturer. An instrumentation purchase choice implies a commitment to a relationship with a particular brand for the useful lifespan of the instrument. It is important for suppliers of confocal instrumentation to remain abreast of cutting edge technology, but it is also important for demonstrate a commitment to regular maintenance and dedication to support of established instruments. Four areas you may wish to get candid information on include:
1.) Availability of service personnel. Service engineers should have effective support with scheduling, and should be able to provide an accurate timeline to the facility visit.
2.) Effective communication between the domestic and global service infrastructure as well as between service management and facility managers.
3.) Parts availability. Replacement parts should be available, and these parts should be ensured to perform acceptably before installation
4.) Preventative maintenance. Heavily used, research critical LSM instrumentation should be overhauled in a disciplined manner at regular intervals to ensure maximum reliability. The rate at which certain components deteriorate should be predictable, and a thorough checklist which ensures that all the components are performing up to specification at regular intervals would do much to bolster end user confidence.
That being said, I feel that the Leica SP-2 showcases some elegant engineering solutions and I'm not infrequently re-impressed with the instrument's capabilities. When the platform is in top working condition I would put it up against any other laser scanning microscope (the newer AOBS feature seems to be a promising development as well, but we don't presently have that capability). I haven't had nearly as much experience with the META at this point, but I'm sure there are facilities which would be happy to provide you with input about instrument performance.
Good luck in your decision--feel free to contact me directly if you have specific questions or concerns.
Best Regards, Karl G.
On Saturday, February 1, 2003, at 01:27 PM, Pitts, Betsey wrote:
} ----------------------------------------------------------------------- } - } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ----------------------------------------------------------------------- } . } } } } Hi- } we are in the process of purchasing a new confocal, and are looking } at a Zeiss510 Meta or a Leica AOBS or SP2. I think that in the demo } process } we are seeing what we need to, and are impressed by the capabilities of } each. I would appreciate user perspectives that we cannot get from the } manufacturers though- are there instrument aspects of either } microscope that } are repeatedly problematic or limiting? Do multi-user facilities have } particular difficulty with either? I do not expect an obvious } "winner" for } an answer, but am more interested in hearing what some of the ups and } down } are, and seeing if those problems or circumstances would be issues for } us. } Thanks for any experience you an offer, I am sure it will be useful. } } Betsey } *********************************************************************** } * } Betsey Pitts } Research Associate/Facilities Manager, Microscopy } The Center for Biofilm Engineering } } 366 EPS Building, Montana State University } Bozeman, MT 59717 } } betsey_p-at-erc.montana.edu } Ph (406) 994-7813 } Fax (406) 994-6098 } } *********************************************************************** } * } } 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, Champaign 61801 Office: B650J Phone: 217-244-6292 Fax: 217-244-6219
Hello Everyone I may be opening a can of worms here but....
What do TEM labs do about the dust created when polymerised resin is cut with a hacksaw or rasp or razor blade. Any resin - Spurr's, Epon, HM20, LR Gold, LR White, etc etc.
Is there a vaccum system recommended? Or do you just use a wet towel system. Or brush the dust into the garbage can and create dust in the atmosphere and not worry about it. Do you have a policy of only cutting resin down to manageable size in a fume hood and then vaccum up the dust? Elaine
-- Dr. Elaine Humphrey Director, BioImaging Facility First Vice 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
Hi All: I am looking for online responses as suggestions of KNOWN suppliers of Denka M3 Lab6 filaments for a JEOL TEM. I would invite offline responses as to specific prices. (Essentially, I am comparison shopping for the supplier with the best prices). Thanks in advance, Michael Coviello Lab Manager, UT Arlington
Hello Everyone The annual meeting of the Microscopical Society of Canada is meeting in Vancouver this year June 4-6, 2003 at the University of British Columbia.
This is a call for papers for two concurrent session in the Biological and Physical Sciences
Instructions for authors, registration and accomodation can be found on the website http://www.emlab.ubc.ca
The list of workshops, exhibitors, local University tours, and invited speakers should be on the website soon. There are links to Tourism Vancouver should you wish to extend your visit to one of the most beautiful places on this planet. (You can probably tell - I am biased).
And there are links to the International Cryo EM course which will be in the following week. Elaine
-- Dr. Elaine Humphrey Director, BioImaging Facility First Vice 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
} Is there a vaccum system recommended? Or do you just use a wet towel } system. Or brush the dust into the garbage can and create dust in the } atmosphere and not worry about it. Do you have a policy of only } cutting resin down to manageable size in a fume hood and then vaccum } up the dust?
I personally use the wet paper towel system. Wrap it up well and then throw it in the trash. Of course, then I wonder about its ultimate fate. Over here a large proportion of our wastes go to an incinerator to generate electrical power, and then I worry about toxic vapors released into the atmosphere. Can't decide if that's better or worse than landfill, where stuff can leach into our water system!
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
Sergey is correct, disk fragmentation has little effect on file size. If you have the program preferences set to "allow fast saves" MS Word will save the recent changes made to your file, along with the original file, thereby making the total file size ever larger (until a certain size is reached after which the file is saved as an original). If you choose "Save as" then the document is saved as an original, minimizing the file size. You can force the smallest size files by de-selecting "allow fast saves"; it will take only a fraction of a second longer to save each time (unless you are writing a book and have a huge file).
Kim {} {} {} {} {} Kim Rensing PhD Department of Botany, UBC 6270 University Blvd. Vancouver BC, Canada V6T 1Z4
On Friday, January 31, 2003, at 08:43 PM, Sergey Ryazantsev wrote: } Gary } I think, you wrong: fragmented file occupied more physical space on HD } (yes) but fragmented/non-fragmented files has the same bit-size. So } fragmentation may affect the reading/writing time only, which on the } modern computers is negligibly. If the disk seriously fragmented, } yes, it may affect computer's performance in general. "Save as" } command not necessary writes in non-fragmented space. NTFS usually } decently manage this issue and keep fragmentation at relatively low } level, but it's OS decision where to place your file (for instance, if } file is small, it would be placed in the NTFS analog of FAT at the } beginning of drive- this is special precaution against fragmentation). } If the HD is seriously fragmented, even "Save as" will cause the file } fragmentation. Sergey } } At 08:18 AM 1/31/2003, you wrote: } } } } What usually happens is that the document's file } } becomes fragmented. In this case, more sectors } } on disk are used (wasted) and leads to a larger } } file size. If you routinely keep disk fragmentation } } low, then do a Save As with the same file name. } } This should put the new save in a contiguous } } area. } } } } gary g } }
Hello all, first off, I am a complete novice, last time I looked through a microscope was in high school, too many years ago to mention. I have done some research though-so I'm not completely in the dark. (Besides, I'm a geek in disguise anyway.) What I want to do is to be able to examine samples of my sourdough starter to see if, and if possible, what kinds of lactobacillis are present in the starter. I have a scope w/capability of 970 power (10x/97xOil)... I was able to stain a sample of yogurt w/methylene blue sucessfully and see some rod shaped bacteria in that slide. They were smaller than I thought they would be, but I could distinguish them. My attempts w/the sourdough are less sucessful. I take a sample of the starter and dilute it and then put a drop on the slide, dry it, pass it through the flame, and stain it. I've found instructions for preparing slide with crystal violet that are a little more involved, using counterstaining etc. Would I be better off getting some cyrstal violet and using this procedure, rather than the more simple method w/the methylene blue, or am I just trying to do something that isn't really possible w/a scope w/only 970 capability. Any advice or direction to documents that would help would be preciated. -cj-
Hello Elaine Nice to hear from you. I was on your Cryo-EM course in the summer last year and still not received yet the samples, which supposed to be processed on that course. If I do remember correctly you promised to me to sent those samples ASAP. Is it still possible to get them back? Your course was very costly to me. The samples were (it was discussed with you prior I signed for the course) from trangenic mice with BM transplantation. The cost of such mice is a few thousand $$ each... As a matter of fact I was disappointed how the Cryo-course where handled. The samples were not processed in time (was sitting in refrigerator waiting for what?), specific antibodies were not ordered in time and then where lost, we did not have necessary reagents in time and equipment was constantly busy... I understand, it's normal laboratory life when you have to make reagent at the last moment (in the cosy environment of your own Lab, not in yours) or wait for equipment available, but it was Cryo-course I paid for that nearly from my pocket a few thousand $$. I was expecting to have at least attention to my needs at such price. I am sorry I made this public statement, but it seems to me, people should know what they may expect to have from the course and I am still WANT BACK MY SAMPLES! Sergey
At 03:05 PM 2/3/2003, 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 Box 951737 Los Angeles, CA 90095-1737
Because of a recent posting I find it appropriate to remind you all of the Listserver Rules, which you all received copies of upon subscription confirmation. In particuliar I draw your attention to #4. If you have a problem with an organization, this is NOT the place to air it.
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Nestor just informed me that my posting (see below) is inappropriate on this ListServer, because I violate the rule. Rule #4 states:
# 4.) This forum is not to be used as a platform to accuse or defame any individual or organization in any negative manner. You may disagree with any comment posted and post a reply, but this server may not be used to spread misleading, derogatory or disparaaging comments under any conditions.
I am deeply apologize, I broke the rule and let you to be exposed to my message. I did not intend to hurt anybody, but express my sincere impression about mentioned here CryoEM course. I also deeply concerned I could not receive my very valuable samples for more than 6 month. Again, I apologize, I posted my message against the rules. I already contact to Nestor, so he could help me to understand what was wrong in my message and how I could avoid mistakes in the future. Sincerely, Sergey Ryazantsev.
At 08:38 PM 2/3/2003, 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 Box 951737 Los Angeles, CA 90095-1737
Hi Folks, Is there any one out there who knows how to refurbish windows for gas flow counters? These are for a JEOL 733. I've a heap of mylar and a stack of old windows and no idea what to do. I would like to know, how to clean the old stuff off, how thick the mylar should be, how to attach a new bit and whether these need C coating afterwards and if so how thick a layer. Cheers, Malc -- Dr MP Roberts Phone: [+27](0)46 603 8313 (work) Dept of Geology [+27] (0)46 6361197 (home) Rhodes University Fax: [+27](0)46 622 9715 6140 Grahamstown Cell: 083 4060 262 SOUTH AFRICA e-mail: m.roberts-at-ru.ac.za
I always try to use an appropriate size of embedding mold to minimise the need for cutting a lot of resin and make sure that the specimen is as close to the tip as possible. Trimming should then be possible by razor blade or glass knife on the microtome and I haven't used a hacksaw on a resin block for some time.
In the unlikely event that I need to saw a specimen I would do so in the fume hood using a small detachable vice to hold the block. The dust would be collected using a damp paper towel, placed in a pot and then dried. This can easily be made safe later by topping up with some waste unpolymerised resin and polymerising for disposal as normal waste.
Many years ago we used an ordinary portable vacuum cleaner but very quickly realized that this would churn out the most hazardous particle sizes of dust. There are apparently lots of new vacuum cleaners with filters in them but I don't know whether their performance would be guaranteed for potentially carcinogenic dust. I have, however, seen a couple of portable machines advertised for disposal of photocopier toner dust which may be suitable. I still don't think that resin dust should be encouraged even if you intend to clean it up later because you don't know how much is already airborne.
I have little worry about the slivers and chips of resin created by razor and glass knife cutting because they are too big to be inhaled (as far as I know), settle very quickly and can be brushed into a waste pot easily. But I would be very concerned about brushing, blowing or vacuuming ~ micron size particles. Of course 20+ years ago information was a bit sparce about the hazards of resins and their dusts and we treated them with a lot less care.
Good luck
Malcolm
Malcolm Haswell e.m. unit School of Sciences University of Sunderland UK ----- Original Message ----- } From: Elaine Humphrey {ech-at-interchange.ubc.ca}
Thanks everybody, for your responses. we finally found water in the HT tank. Trying to arrange for oil to change it.
Shashi Singh Scientist CCMB, Hyderabad INDIA
===== Shashi Singh Scientist Centre for Cellular and Molecular Biology Hyderabad-500 007 INDIA PH-91-40-7192575,7192761,7192615 FAX-91-40-7160591, 7160311
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The program committee for M&M 2003 would like to remind everyone that presentation submissions are due in less than two weeks (February 17). Instructions for submission and details about the meeting are available on-line at:
The meeting is shaping up to be very exciting with particular focus on Nanotechnology and Optical Microscopy, as well as the broad coverage of Electron Microscopy techniques and applications that you expect at M&M. The program committee encourages you to participate by submitting a paper on your work, and we hope to see you in San Antonio!
I just want to say that the recent comment by Sergey to Elaine Humphrey re: problems with the Cryo course at last year's MSC meeting was inappropriate. He has a personal complaint that should have been directed specifically to Elaine. It was totally uncalled for to air it to the whole ListServer community.
I've noticed that this type of grudge comments do occur occasionally on the ListServer and take up valuable time and space. Please limit comments like that to the person involved.
Peggy Sherwood
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
In our lab we have an plain old household vaccum cleaner that is used to suck up the dust. The hose used to be set up on a stand so that it could suck up the dust as it is made, but the stand wasn`t functional when I got here 2 years ago, so I couldn`t tell you how it worked. These days I just vaccum the dust up as I go along and try not to let it get all over the place (we don`t work in a fume hood). The vaccum bag has never needed to be emptied in my time here, so I have no idea how the waste is dealt with.
I hope this helps, Robin __________________________ Robin Elizabeth Young Laboratoire de Jacques Paiement Université de Montréal re.young-at-umontreal.ca
} What do TEM labs do about the dust created when polymerised resin is } cut with a hacksaw or rasp or razor blade. Any resin - Spurr's, Epon, } HM20, LR Gold, LR White, etc etc. } } Is there a vaccum system recommended? Or do you just use a wet towel } system. Or brush the dust into the garbage can and create dust in the } atmosphere and not worry about it. Do you have a policy of only } cutting resin down to manageable size in a fume hood and then vaccum } up the dust?
Elaine and Robin: If you are using a plain old vacuum, I would bet you are capturing the big particles and simply exhausting the small, more dangerous ones. that's why a regular vacuum is worse for some allergic to dust. They make vacuums with HEPA filters now but I don't know if they are really effective. the water trap ones are not according to studies i have read. the best option for vacuums is one that vents to the outside (e.g., a built-in whole house vacuum) and these are widely recommended for those with bad dust allegies. The bottom line is that you may be making things worse since you are distributing them into the air.
We use a Dremel moto-tool for trimming our blocks. Vastly superior to a hacksaw or razor blade in my opinion but it generates a ton of dust. I would never do it outside the fume hood. My guess is you would be risking the equivalent of silicosis. Tom
At 10:49 AM 2/4/2003 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Associate Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
Dear Malc, It has been a long time since I did this on my JEOL JXA-3A, but we used to use 4 or 6 micron mylar, stretch it over the window and glue it on with 5-minute epoxy, then trim off the excess. I believe we sanded off the old epoxy from the brass window holder. The windows are usually then lightly aluminum-coated so they won't charge. Good luck Mary Mager Electron Microscopist Metals and Materials Eng. UBC 6350 Stores Rd. Vancouver, B.C. CANADA tel: 604-822-5648 fax: 604-822-3619 ----- Original Message ----- } From: "Malc" {m.roberts-at-ru.ac.za} To: "Microscopy discussion group" {Microscopy-at-sparc5.microscopy.com} Sent: Tuesday, February 04, 2003 1:57 AM
The answer is that Alexa 647 has a component that, when excited at 470 to 490 nm, has an emission very similar to FITC. This has been confirmed by our spectrophotometer, a colleague with the Zeiss Meta (posted on the list) and Molecular Probes themselves.
Practically, this means that while the dye works great at 647 nm, it cannot be used with GFP or other staining in the green range.
We're gonna try Alexa 633.
Thanks!! ____________________________________________________________________________ Michael Cammer Analytical Imaging Facility Albert Einstein Coll. of Med. Jack & Pearl Resnick Campus 1300 Morris Park Ave. Bronx, NY 10461 (718) 430-2890 Fax: 430-8996 URL: http://www.aecom.yu.edu/aif/
If there is texture on the surface, what it the general size of the features? Also, what is the thickness specification for the coating?
If you want to do thickness measurements non-destructively, have you considered Raman confocal?
There are a number of approaches to this problem. Please feel free to call me off-line for further discussion.
Best regards, Barbara Foster Microscopy/Microscopy Education 125 Paridon Street, Suite 102 Springfield, MA 01118 PH: 413-746-6931 FX: 413-746-9311 Web: www.MicroscopyEducation.com
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At 05:21 PM 1/31/03 -0600, Chris Michaelson (by way of wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} } The Al coatings on thin window detectors do a good job of } keeping out visible light, but are fairly transparent to IR. } The IR photons are absorbed just like X-rays, but because } their energy is so low and the flux high compared to X-ray } photons, the result is a nearly continuous flow of current } through the detector which is indistinguishable (to the } pulse processing electronics) from a "leaky detector".
Just off-the-cuff, and in ignorance, I'm surprised that IR has enough energy to produce any elextrons/holes at all, as I didn't think it would get through the Au layer.
I guess this must be the right explanation, though
cheers
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
Has anyone tried the Leica EM Trim specimen trimmer?
Uses a milling tool to trim down blocks, while observing througha stereo microscope head. Has a vacuum connection to trim away the nasties.
Thinking of buying one so any feed back would be appreciated.
Allan -- ------------------------------------------------- Allan Mitchell Technical Manager Otago Centre for Electron Microscopy C/-Department of Anatomy and Structural Biology School of Medical Sciences P.O. Box 913 Dunedin New Zealand
I want to thank all who responded with offers for manuals. Someone was able send me a scanned copy that is as close to an original as I could hope for. I understand that the arm bearing surfaces are probably damaged. The instrument was transported without using the shipping blocks, even though they had them in the drawers. So my question is, does anyone know of a source for replacement specimen arm bearings? I already called Leica and they said good luck. Very best regards to all who replied, Steve D'Angelo
--
Equipment Resurrection 1005 Terra Nova Boulevard, Suite 2 Pacifica, CA 94044. 650-738-0351 http://equiprx.net/
I bought a hand held domestic vacuum cleaner. Then one day I was using it over a black bag and noticed that it appeared to be depositing fine dust.
I returned to the old method. I do sawing inside a large plastic bag in a fume hood. I knot the bag and store it the cupboard (which is vented by the system) under the hood. When it is full in oh... 2024 when I retire I will polymerise it.
Dave
On Tue, 04 Feb 2003 10:42:28 -0600 Tom Phillips {phillipst-at-missouri.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Elaine and Robin: If you are using a plain old vacuum, I would bet you are } capturing the big particles and simply exhausting the small, more dangerous } ones. that's why a regular vacuum is worse for some allergic to } dust. They make vacuums with HEPA filters now but I don't know if they are } really effective. the water trap ones are not according to studies i have } read. the best option for vacuums is one that vents to the outside (e.g., a } built-in whole house vacuum) and these are widely recommended for those } with bad dust allegies. The bottom line is that you may be making things } worse since you are distributing them into the air. } } We use a Dremel moto-tool for trimming our blocks. Vastly superior to a } hacksaw or razor blade in my opinion but it generates a ton of dust. I } would never do it outside the fume hood. My guess is you would be risking } the equivalent of silicosis. Tom } } } } At 10:49 AM 2/4/2003 -0500, you wrote: } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } Hi Elaine, } } } } In our lab we have an plain old household vaccum cleaner that is used to } } suck up the dust. The hose used to be set up on a stand so that it could } } suck up the dust as it is made, but the stand wasn`t functional when I got } } here 2 years ago, so I couldn`t tell you how it worked. These days I just } } vaccum the dust up as I go along and try not to let it get all over the } } place (we don`t work in a fume hood). The vaccum bag has never needed to be } } emptied in my time here, so I have no idea how the waste is dealt with. } } } } I hope this helps, } } Robin } } __________________________ } } Robin Elizabeth Young } } Laboratoire de Jacques Paiement } } Université de Montréal } } re.young-at-umontreal.ca } } } } } What do TEM labs do about the dust created when polymerised resin is } } } cut with a hacksaw or rasp or razor blade. Any resin - Spurr's, Epon, } } } HM20, LR Gold, LR White, etc etc. } } } } } } Is there a vaccum system recommended? Or do you just use a wet towel } } } system. Or brush the dust into the garbage can and create dust in the } } } atmosphere and not worry about it. Do you have a policy of only } } } cutting resin down to manageable size in a fume hood and then vaccum } } } up the dust? } } Thomas E. Phillips, PhD } Associate 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 } } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
Steve: I don't know where you are located, but Helmut Patzig, of MOC, Valley Cottage, NY, keeps the OMU-3 here running with the odd bits. Phone number is 845-268- 6450. He might have the arm or bearing (I damaged one of those many, many, many years ago!!) and some other bits you will likely need. However, _no one_ seems to have any more of the drive belts!
Roger Moretz, Ph.D. Dept of Toxicology BI Pharmaceuticals -- Where the world is only slightly less weird than it actually is. } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } I want to thank all who responded with offers for manuals. } Someone was able send me a scanned copy that is as close to an original } as I could hope for. } I understand that the arm bearing surfaces are probably damaged. } The instrument was transported without using the shipping blocks, even } though they had them in the drawers. } So my question is, does anyone know of a source for replacement specimen } arm bearings? } I already called Leica and they said good luck. } Very best regards to all who replied, } Steve D'Angelo } } -- } } } Equipment Resurrection } 1005 Terra Nova Boulevard, Suite 2 } Pacifica, CA 94044. } 650-738-0351 } http://equiprx.net/ } } }
I have just seen a note in Microscopy Today, (downloaded from http://www.microscopy-today.com follow the links to the back issue Table of Contents), (March/April 2002).
Karen Pawlowski uses a method that traps the dust in water in a flask.
Dave
On Tue, 04 Feb 2003 10:42:28 -0600 Tom Phillips {phillipst-at-missouri.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Elaine and Robin: If you are using a plain old vacuum, I would bet you are } capturing the big particles and simply exhausting the small, more dangerous } ones. that's why a regular vacuum is worse for some allergic to } dust. They make vacuums with HEPA filters now but I don't know if they are } really effective. the water trap ones are not according to studies i have } read. the best option for vacuums is one that vents to the outside (e.g., a } built-in whole house vacuum) and these are widely recommended for those } with bad dust allegies. The bottom line is that you may be making things } worse since you are distributing them into the air. } } We use a Dremel moto-tool for trimming our blocks. Vastly superior to a } hacksaw or razor blade in my opinion but it generates a ton of dust. I } would never do it outside the fume hood. My guess is you would be risking } the equivalent of silicosis. Tom } } } } At 10:49 AM 2/4/2003 -0500, you wrote: } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } Hi Elaine, } } } } In our lab we have an plain old household vaccum cleaner that is used to } } suck up the dust. The hose used to be set up on a stand so that it could } } suck up the dust as it is made, but the stand wasn`t functional when I got } } here 2 years ago, so I couldn`t tell you how it worked. These days I just } } vaccum the dust up as I go along and try not to let it get all over the } } place (we don`t work in a fume hood). The vaccum bag has never needed to be } } emptied in my time here, so I have no idea how the waste is dealt with. } } } } I hope this helps, } } Robin } } __________________________ } } Robin Elizabeth Young } } Laboratoire de Jacques Paiement } } Université de Montréal } } re.young-at-umontreal.ca } } } } } What do TEM labs do about the dust created when polymerised resin is } } } cut with a hacksaw or rasp or razor blade. Any resin - Spurr's, Epon, } } } HM20, LR Gold, LR White, etc etc. } } } } } } Is there a vaccum system recommended? Or do you just use a wet towel } } } system. Or brush the dust into the garbage can and create dust in the } } } atmosphere and not worry about it. Do you have a policy of only } } } cutting resin down to manageable size in a fume hood and then vaccum } } } up the dust? } } Thomas E. Phillips, PhD } Associate 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 } } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
I hate the way Word deals with images. It makes huge files too. I would recommend Adobe Pagemaker or Deneba Canvas for much more controllable combination or text and graphics. If you have Acrobat (full version) installed you can then also export the files to pdf format and share them easily on the web. Of course, these software packages are not so ideal for presentations and I have not yet found an alternative to Powerpoint for that.
(P.S. This is not just Microsoft bashing, I wish they would do a better job of Graphics handling in word. In my opinion, the competition is better at this just now. As a pure Word processor I find MS Word very good).
Best wishes
Ian
On Mon, 20 Jan 2003 09:30:47 -0600, {"gary.m.brown-at-exxonmobil.com"-at-sparc5.microscopy.com} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } Jeff, } } I, too, have had significant problems working with images imported into } Microsoft Word. However, my software is located on the PC hard drive. The } biggest problem that I have encountered with images in Microsoft Word } occurs when annotating images. After the image is imported into Word, } annotation may be done in two ways (to my knowledge): (1) Text, arrows, } etc. may be simply superimposed over the images. The problem with this } approach is that the annotations are not linked to the image and may not } remain superimposed on the image if the image moves. (2) Annotations can } also be linked (probably not the best choice of words) to the image by } double-clicking on the image to open the image field, adding the } annotation, then closing the image field. These annotations are permanent } unless intentionally moved or deleted. } } The problems occur when one implements the second option. Comparing } images } before and after annotation, I found that the annotated images often } sustained substantial changes in gray or color levels. Case in point, EDS } maps were so badly affected that the color key was no longer correct. } } My solutions follow: (1) Annotate images in Adobe Photoshop before } importing into Word. Note that the effects of lossy compression on } annotations (blurred edges) may be pronounced. (2) Use Microsoft } PowerPoint } for image presentation. I have encountered no problems with image files } in } PowerPoint. } } Good luck to you in your endeavors. } } Cheers, } } "The opinions expressed are those of Gary M. Brown and do not represent } the } opinions of ExxonMobil Corporation nor its affiliates." } } Gary M. Brown } ExxonMobil Chemical Company } Baytown Polymers Center } 5200 Bayway Drive } Baytown, Texas 77520-2101 } phone: (281) 834-2387 } fax: (281) 834-2395 } e-mail: Gary.M.Brown-at-ExxonMobil.com } } } "Oakley, Jeff" } {oakleyj-at-rayovac.c To: {Microscopy-at-sparc5.microscopy.com} } om} cc: } Subject: RE: presentation images } 01/17/03 07:50 AM } } } } } } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } This same phenomenon happens with my reports in Microsoft Word. In } addition to images darkening, they sometimes shift to other pages and/or } change size and dimension. Adding tables and text boxes to the report } adds } to the fun. } } The software that we use is networked. Our IS department has told us } that } the networked software has a bug that causes these things to happen when } file sizes increase, and that there is not a patch for it. So we have } just } have to deal with it. } } Jeff Oakley } } } -----Original Message----- } } From: Corazon D. Bucana [mailto:bucana-at-audumla.mdacc.tmc.edu] } Sent: Thursday, January 16, 2003 11:11 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: presentation images } } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } I created a Power point presentation last November 2002 consisting of } several fluorescence micrographs. The file which was rather large ( 90 } Mb) } } was left in my laptop all this time. When I opened it again this week I } find that the images are now too dark and I needed to increase brightness } by 3-4 clicks on the brightness icon of Power point.I increased } brightness } } of all the micrographs and copied the file to a CD hoping that the image } will not deteriorate there and then compare it with the one in my laptop } several weeks from now. Has this happened to anyone else? Is there } something I should have done to prevent this? } } Any suggestions or comments will be greatly appreciated. } } Cora Bucana } } } } } } } } }
-- Ian MacLaren Technische Universität Darmstadt Material-und Geowissenschaften Petersenstr. 23 64287 Darmstadt Germany
I'm curious as to how many people have had problems using molecular sieves in dehydration solvents, with respect to knife damage. We seem to be going through diamond knives at a uncomfortable rate and we're wondering if this could be a contributing factor. We are very careful with our knives and try to minimize contact cleaning of the edges. We soak the knives often in the recommended solution in a commercial cleansing unit, and the knife manufacturer has evaluated one of our knives and confirmed that the edge is chipped, not dirty.
As a multi-user facility, we do a LOT of ultramicrotomy on a large variety of samples, so this may just be normal wear and tear, but we would sure like to minimize this expense.
Thanks much!
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We're the Fun Core! 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/
I was wondering if anyone on the list is familiar with this stain. I am trying to find a procedure for either making this stain or a commercial source for the stain. It was listed in a methods section for negative staining isolated neurofibrillary tangles (Crowther, R. A. 1991. PNAS 88:2288) but the author did not reference a source for the stain or a procedure. I would appreciate any help provided.
Thanks
Lawrence X. Oakford, Ph. D. Technical Manager Microscopy Core Facility Department of Cell Biology and Genetics UNT Health Science Center 3500 Camp Bowie Blvd. Fort Worth, TX 76107
Preparing my abstract for MM'03 I have found that: - Two page document (4 images with superimposed EDS line scans) saved in Word 2000 format was 758K size. - Saved in Word 6.0 format (as required for uploading by submission instructions) it had size of 2.93M (!). - Saved in PDF format (Acrobat 5.0) file was just 286K, but line scans, still pretty visible, were looking not nice.
I believe I should upload PDF files, since Word files anyway will be converted in PDF for CD publishing. But are not we loosing quality going digital now?
Vladimir
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: Rosemary White [mailto:rosemary.white-at-csiro.au] } Sent: Thursday, January 30, 2003 4:00 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Re: presentation images in Microsoft Word } } } -------------------------------------------------------------- } ---------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ---------. } } } Another trick is to put the images and text into a Word } table, if you size } the cells in the table to about the size you want, the images } will insert } to fit the cell size. } } And if copying and pasting, make sure you do "Paste Special" } and unclick } the box that says "float over text". That way the images } always stay in } the same spot with respect to the surrounding text and don't } mysteriously } jump around when you insert or delete text. } cheers, } Rosemary } } } } Thanks Doug- } } Great tips- I'm printing them up until I memorize them. } } Rgds, } } Mike Shaw } } Roselle, NJ } } } } } Gary, } } } } } } MS Word is such a pain because of the way it works with } images. A couple } } } of tricks I've discovered over the years. } } } } } } 1) Place the image within a text box, rather than directly } on the page, it } } } seems to give you much greater control over the location } on the page. It } } } also makes it much easier to add text annotations that } stay with the image. } } } } } } } } } If you don't want the text box to have a border you can } remove it by } } } selecting } } } the box outline and look for the "paint brush" icon on the } Draw toolbar, } } } then use the down arrow and select "none". If you'd like } the text box to } } } be transparent, select the text box outline and look for } the "paint bucket" } } } icon on the Draw toolbar, then use the down arrow and } select "none". If } } } you discover its hard to find the text box border once you } made the edge } } } "invisible", first select the image with a single left } click (it should have } } } the solid black resizing "handles") and then use the } keyboard left or right } } } arrow keys and the selection with move out to the textbox } outline (with } } } black } } } bordered resizing "handles"). } } } } } } 2) Always use the INSERT | PICTURE | FROM FILE option as } opposed to copying } } } and pasting an image into Word. I find that the images } are harder to work } } } with if I paste them in. Also, you can insert TIFF or BMP } } } images into Word, } } } you don't have to use JPEG. } } } } }
I run a clinical EM lab with a geriatric JEOL 100C TEMSCAN microscope. I am campaigning for new equipment and would like to get a stand alone TEM and SEM. Currently my scope is in a rather large room that should be able to accommodate two instruments with their columns at least ten feet apart. I would hate to have new instruments installed only to find that they interfered with each other, either electrically or mechanically (vibration, etc.). I would like to hear from anyone who has wrestled with this problem....or is it a problem? I am on the second floor and not too near elevator shafts or electrical traces and vibration for my one scope has not been a problem.
Joiner Joiner Cartwright, Jr., Ph.D. Department of Pathology Baylor College of Medicine Houston, Texas U.S.A.
The New England Society for Microscopy announces its Early Spring meeting, to be held at the JEOL(USA) Inc. facility in Peabody, Massachusetts.
This is something we discovered over 20 years ago: molecular sieves work well BUT it is necessary to allow the alcohols to stand untouched for one month in order for the ceramic-like "fines" to settle out. Also, be very careful when withdrawing sieve-dried alcohols. Do not pour the alcohols but use a pipette and remove it from the top of the liquid. When the level drops to less than 1 inch above the sieves, it's time to move on to the next bottle. Basically, we would prepare about 6-10 pints of ethanol at one time, allowing them to "age" for at at least 30 days.
I must admit that now I tend to use 100% ethanol right out of freshly opened containers (individually sealed pints) except in the most critical of applications (Spurr's dehydrations, for example) and have NEVER had a problem with water.
I know of at least two investigators (not me) who have damaged diamond knives by not taking precautions with molecular sieves. Basically, once the fines get onto your specimen they are impossible to remove and will damage your diamond knife.
} I'm curious as to how many people have had problems using molecular } sieves in dehydration solvents, with respect to knife damage. We seem } to be going through diamond knives at a uncomfortable rate and we're } wondering if this could be a contributing factor. We are very careful } with our knives and try to minimize contact cleaning of the edges. We } soak the knives often in the recommended solution in a commercial } cleansing unit, and the knife manufacturer has evaluated one of our } knives and confirmed that the edge is chipped, not dirty. } } As a multi-user facility, we do a LOT of ultramicrotomy on a large } variety of samples, so this may just be normal wear and tear, but we } would sure like to minimize this expense.
############################################################## John J. Bozzola, Ph.D., Director I.M.A.G.E. (Integrated Microscopy & Graphics Expertise) 750 Communications Drive - MC 4402 Southern Illinois University Carbondale, IL 62901 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu ##############################################################
Thank you, Valerie. That's a good point about monitor light, etc. and perhaps a curtain arrangement might help. As for the number of people in the confined space....Is this an advantage or disadvantage??
Joiner ++++++++++++++++++++++++++++++ At 03:04 PM 02/06/2003 -0500, you wrote: We have two microscopes (a JEOL 100S TEM and a Philips 505 SEM) within a space of about 15 X 20 feet, but separated into 2 rooms, each about 15 X 10 ft. The columns are about 11 feet apart and we have absolutely no problems with interference or vibration. However, unless both of the new microscopes can be operated in room light (i.e. have computer monitors & digital capture), putting both scopes into one room could be a problem when both scopes need to be used at the same time. Also, if the space is really small, do you want that many people in there at one time? Having space for the equipment is one thing, but room for the users too, is another.....
Valerie
+++++++++++++++++++++++++++++++
Hello Listrers -
I run a clinical EM lab with a geriatric JEOL 100C TEMSCAN microscope. I am campaigning for new equipment and would like to get a stand alone TEM and SEM. Currently my scope is in a rather large room that should be able to accommodate two instruments with their columns at least ten feet apart. I would hate to have new instruments installed only to find that they interfered with each other, either electrically or mechanically (vibration, etc.). I would like to hear from anyone who has wrestled with this problem....or is it a problem? I am on the second floor and not too near elevator shafts or electrical traces and vibration for my one scope has not been a problem.
Joiner Joiner Cartwright, Jr., Ph.D. Department of Pathology Baylor College of Medicine Houston, Texas U.S.A.
Our lab purchased a Leica EM Trim about a year ago. It does a pretty good job of trimming. The vacuum cleaner (purchased separately) comes on automatically when you push the trimming button and does a good job of keeping the dust down.
Gene P. Young Sr. Analytical Technologist Analytical Sciences, Polymer Characterization The Dow Chemical Company 2301 N. Brazosport Blvd., B-1470 Freeport, Texas 77541-3257 Fax: (979) 238-0095 Phone:(979) 238-1579
-----Original Message----- } From: Allan Mitchell [mailto:allan.mitchell-at-stonebow.otago.ac.nz] Sent: Tuesday, February 04, 2003 4:05 PM To: Microscopy-at-sparc5.microscopy.com
Has anyone tried the Leica EM Trim specimen trimmer?
Uses a milling tool to trim down blocks, while observing througha stereo microscope head. Has a vacuum connection to trim away the nasties.
Thinking of buying one so any feed back would be appreciated.
Allan -- ------------------------------------------------- Allan Mitchell Technical Manager Otago Centre for Electron Microscopy C/-Department of Anatomy and Structural Biology School of Medical Sciences P.O. Box 913 Dunedin New Zealand
You can set the resolution of your images higher in Adobe Acrobat. I don't know which Acrobat version you have and it is a little different in each. The easiest way is to use PDFWriter and set the resolution output to 300 dpi or a little higher. If you use Distiller, then you will need to go in and change the resolution of your gray scale and line drawings to higher and also make sure that you are using a higher resolution print option. You can set everything to 300 dpi and it should come out pretty good. You can also set your line drawing settings higher than 300 if you want.
In adobe Acrobat 5.0, I save in 4.0 format and for documents that I save for myself for reference, I use a general resolution of 600 dpi. For image compression schemes, I use bi-cubic sampling to 300 dpi for images above 350 dpi for both color and grayscale, and 600 for monochrome images. These settings work fairly well for me and the document sizes don't get too large.
-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: Dusevich, Vladimir [mailto:dusevichv-at-umkc.edu] Sent: Thursday, February 06, 2003 11:25 AM To: Microscopy-at-sparc5.microscopy.com
Hi Listers,
Preparing my abstract for MM'03 I have found that: - Two page document (4 images with superimposed EDS line scans) saved in Word 2000 format was 758K size. - Saved in Word 6.0 format (as required for uploading by submission instructions) it had size of 2.93M (!). - Saved in PDF format (Acrobat 5.0) file was just 286K, but line scans, still pretty visible, were looking not nice.
I believe I should upload PDF files, since Word files anyway will be converted in PDF for CD publishing. But are not we loosing quality going digital now?
Vladimir
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: Rosemary White [mailto:rosemary.white-at-csiro.au] } Sent: Thursday, January 30, 2003 4:00 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Re: presentation images in Microsoft Word } } } -------------------------------------------------------------- } ---------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ---------. } } } Another trick is to put the images and text into a Word } table, if you size } the cells in the table to about the size you want, the images } will insert } to fit the cell size. } } And if copying and pasting, make sure you do "Paste Special" } and unclick } the box that says "float over text". That way the images } always stay in } the same spot with respect to the surrounding text and don't } mysteriously } jump around when you insert or delete text. } cheers, } Rosemary } } } } Thanks Doug- } } Great tips- I'm printing them up until I memorize them. } } Rgds, } } Mike Shaw } } Roselle, NJ } } } } } Gary, } } } } } } MS Word is such a pain because of the way it works with } images. A couple } } } of tricks I've discovered over the years. } } } } } } 1) Place the image within a text box, rather than directly } on the page, it } } } seems to give you much greater control over the location } on the page. It } } } also makes it much easier to add text annotations that } stay with the image. } } } } } } } } } If you don't want the text box to have a border you can } remove it by } } } selecting } } } the box outline and look for the "paint brush" icon on the } Draw toolbar, } } } then use the down arrow and select "none". If you'd like } the text box to } } } be transparent, select the text box outline and look for } the "paint bucket" } } } icon on the Draw toolbar, then use the down arrow and } select "none". If } } } you discover its hard to find the text box border once you } made the edge } } } "invisible", first select the image with a single left } click (it should have } } } the solid black resizing "handles") and then use the } keyboard left or right } } } arrow keys and the selection with move out to the textbox } outline (with } } } black } } } bordered resizing "handles"). } } } } } } 2) Always use the INSERT | PICTURE | FROM FILE option as } opposed to copying } } } and pasting an image into Word. I find that the images } are harder to work } } } with if I paste them in. Also, you can insert TIFF or BMP } } } images into Word, } } } you don't have to use JPEG. } } } } }
Sounds like you have lossy image compression enabled in Acrobat, or possibly low output resolution settings.
Lossless or uncompressed options are available in Acrobat, and should give you the high quality you need for publishing. There are quality "presets" as well; "prepress" is the high quality option if I remember correctly (or just manually reduce compression and increase output resolution).
I believe there was an article with tips on using Acrobat for scientific publishing in a recent issue of "Microscopy Today". Anyone remember what issue it was?
-Kevin ------------------------------------------------ Kevin Frischmann, Laboratory Manager Microscopy & Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
At 10:25 AM 2/6/03 -0600, "Dusevich, Vladimir" {dusevichv-at-umkc.edu} wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Joiner, Large rooms are a rare blessing! Generally the interference is different light requirements for the 2 instruments. I've seen a number of labs where the TEM will have a light-tight curtain suspended from an overhead rail surrounding it. I believe this is a fairly readily available darkroom curtain Also be sure that each room section has its own light switches and one doesn't interfere with the other. The TEM needs the darkness, while the newer computer based SEMs can operate in full light. An older SEM's light sensitivity is more related to each operator than to the instrument itself. Some people can see the images in lighter conditions than others.
If either instrument is going to be used a lot for high mag, then sound and other sources of vibration from the other instrument's user can be a problem. Also, computers related to EDS and image capture should be kept away from either column.
Ken Converse owner Quality Images third party SEM service Delta,PA
Joiner Cartwright, Jr., Ph.D. wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Hello Listrers - } } I run a clinical EM lab with a geriatric JEOL 100C TEMSCAN microscope. } I am campaigning for new equipment and would like to get a stand alone } TEM and SEM. Currently my scope is in a rather large room that should } be able to accommodate two instruments with their columns at least ten } feet apart. I would hate to have new instruments installed only to } find that they interfered with each other, either electrically or } mechanically (vibration, etc.). I would like to hear from anyone who } has wrestled with this problem....or is it a problem? I am on the } second floor and not too near elevator shafts or electrical traces and } vibration for my one scope has not been a problem. } } Joiner } Joiner Cartwright, Jr., Ph.D. } Department of Pathology } Baylor College of Medicine } Houston, Texas U.S.A. } } }
You might want to check the recent archives, as this topic came up not too long ago. It sounds like you will be getting (if successful!) 'workhorse' instruments, which are less sensitive to fields, etc than 'high-end' instruments. The best bet might be to check with potential vendors re column distances and vibrations.
On the practical side, though, the SEM is a normal-to-low light environment, whereas the TEM has to have a low-to-no light environment. I've heard of places where they've tried to get around this with floor-to-ceiling drapes, but light spillage during plate exposure on the TEM will always be an issue. Chatting during simultaneous usage will also be an irritant. I'm sure space is at a premium, however, so why not consider a partition? One advantage of close-together instruments would be sharing a common water chiller, if that is in the budget.
Tom
Dr. Tom Malis Scientist Advisor Natural Resources Canada Govt. of Canada 613-995-7358 malis-at-nrcan.gc.ca
I run a clinical EM lab with a geriatric JEOL 100C TEMSCAN microscope. I am campaigning for new equipment and would like to get a stand alone TEM and SEM. Currently my scope is in a rather large room that should be able to
accommodate two instruments with their columns at least ten feet apart. I would hate to have new instruments installed only to find that they interfered with each other, either electrically or mechanically (vibration, etc.). I would like to hear from anyone who has wrestled with this problem....or is it a problem? I am on the second floor and not too near
elevator shafts or electrical traces and vibration for my one scope has not been a problem.
Joiner Joiner Cartwright, Jr., Ph.D. Department of Pathology Baylor College of Medicine Houston, Texas U.S.A.
Randy, I've not the foggiest as to what molecular sieves are made of. Are there any 'hard bits' in them, like fine ceramic particles? If so, that could be your problem. In many hard materials edge chipping is a reality. However, when sectioning uniformly hard materials like ceramics, the only way they will section is if one has an ultrafine facet on the block, ie the order of several microns. That methodology also has the benefit of reducing the area of damage along the edge, and one can 'walk' along the edge for some time before resharpening is needed. So if there is something hard/tough enough in your sieves to chip the edge, and you are cutting sections of hundreds of microns, you will surely get chipping spread over that length of the edge. The question than becomes, can your clients live with smaller sections?
Tom
-----Original Message----- } From: Tindall, Randy D. To: microscopy-at-sparc5.microscopy.com Sent: 2/6/2003 10:07 AM
Dear Listers,
I'm curious as to how many people have had problems using molecular sieves in dehydration solvents, with respect to knife damage. We seem to be going through diamond knives at a uncomfortable rate and we're wondering if this could be a contributing factor. We are very careful with our knives and try to minimize contact cleaning of the edges. We soak the knives often in the recommended solution in a commercial cleansing unit, and the knife manufacturer has evaluated one of our knives and confirmed that the edge is chipped, not dirty.
As a multi-user facility, we do a LOT of ultramicrotomy on a large variety of samples, so this may just be normal wear and tear, but we would sure like to minimize this expense.
Thanks much!
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We're the Fun Core! 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/
Hello Randy, We use molecular sieves in our 100% ethanol used in dehydration series and in 15 years have never had a problem associated with this practice. We also use only plastic disposable pipettes to avoid any glass splinters in any of our bottles. Do users in your lab share knives? We once had a student in the lab who rapidly destroyed several knives when he got his hands on them. He continually reported that there was something wrong with the knife! We don't know what he did, but we knew who did it, and we took measures to prevent further damage. Dean Abel Biological Sciences 141 BB University of Iowa Iowa City IA 52242-1324
Were the 4 images imported by reference or included in the final document? Big difference.
gary g.
At 08:25 AM 2/6/2003, you wrote:
} Hi Listers, } } Preparing my abstract for MM'03 I have found that: } - Two page document (4 images with superimposed EDS line scans) } saved in Word 2000 format was 758K size. } - Saved in Word 6.0 format (as required for uploading by } submission instructions) it had size of 2.93M (!). } - Saved in PDF format (Acrobat 5.0) file was just 286K, } but line scans, still pretty visible, were looking not } nice. } } I believe I should upload PDF files, since Word files } anyway will be converted in PDF for CD publishing. But are not } we loosing quality going digital now? } } 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 } } } } } -----Original Message----- } } From: Rosemary White [mailto:rosemary.white-at-csiro.au] } } Sent: Thursday, January 30, 2003 4:00 PM } } To: Microscopy-at-sparc5.microscopy.com } } Subject: Re: presentation images in Microsoft Word } } } } } } -------------------------------------------------------------- } } ---------- } } The Microscopy ListServer -- Sponsor: The Microscopy Society } } of America } } To Subscribe/Unsubscribe -- Send Email to } } ListServer-at-MSA.Microscopy.Com } } On-Line Help } } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -------------------------------------------------------------- } } ---------. } } } } } } Another trick is to put the images and text into a Word } } table, if you size } } the cells in the table to about the size you want, the images } } will insert } } to fit the cell size. } } } } And if copying and pasting, make sure you do "Paste Special" } } and unclick } } the box that says "float over text". That way the images } } always stay in } } the same spot with respect to the surrounding text and don't } } mysteriously } } jump around when you insert or delete text. } } cheers, } } Rosemary } } } } } } Thanks Doug- } } } Great tips- I'm printing them up until I memorize them. } } } Rgds, } } } Mike Shaw } } } Roselle, NJ } } } } } } } Gary, } } } } } } } } MS Word is such a pain because of the way it works with } } images. A couple } } } } of tricks I've discovered over the years. } } } } } } } } 1) Place the image within a text box, rather than directly } } on the page, it } } } } seems to give you much greater control over the location } } on the page. It } } } } also makes it much easier to add text annotations that } } stay with the image. } } } } } } } } } } } } If you don't want the text box to have a border you can } } remove it by } } } } selecting } } } } the box outline and look for the "paint brush" icon on the } } Draw toolbar, } } } } then use the down arrow and select "none". If you'd like } } the text box to } } } } be transparent, select the text box outline and look for } } the "paint bucket" } } } } icon on the Draw toolbar, then use the down arrow and } } select "none". If } } } } you discover its hard to find the text box border once you } } made the edge } } } } "invisible", first select the image with a single left } } click (it should have } } } } the solid black resizing "handles") and then use the } } keyboard left or right } } } } arrow keys and the selection with move out to the textbox } } outline (with } } } } black } } } } bordered resizing "handles"). } } } } } } } } 2) Always use the INSERT | PICTURE | FROM FILE option as } } opposed to copying } } } } and pasting an image into Word. I find that the images } } are harder to work } } } } with if I paste them in. Also, you can insert TIFF or BMP } } } } images into Word, } } } } you don't have to use JPEG. } } } } } } } } } }
WE ARE MEMBERS OF A SPECIAL COMMITTEE FOR BUDGET AND PLANNING OF THE NIGERIAN NATIONAL PETROLEUM CORPORATION (NNPC). THIS COMMITTEE IS PRINCIPALLY CONCERNED WITH CONTRACT AWARDS AND APPROVAL. WITH OUR POSITIONS, WE HAVE SUCCESSFULLY SECURED FOR OURSELVES THE SUM OF TWENTY-ONE MILLION, FIVE HUNDRED THOUSAND UNITED STATES DOLLARS(US$21.5M). THIS AMOUNT WAS CAREFULLY MANIPULATED BY OVER-INVOICING OF AN OLD CONTRACT.
BASED ON INFORMATION GATHERED ABOUT YOU, WE BELIEVE YOU WOULD BE IN A POSITION TO HELP US IN TRANSFERRING THIS FUND (US$21.5M) INTO A SAFE ACCOUNT. IT HAS BEEN AGREED THAT THE OWNER OF THE ACCOUNT WILL BE COMPENSATED WITH 20% OF THE REMITTEDFUNDS, WHILE WE KEEP 70%, AND 10% WILL BE SET ASIDE TO OFFSET EXPENSES AND PAYTHE NECESSARY TAXES.
ALL MODALITIES OF THIS TRANSACTION HAVE BEEN WORKED OUT AND ONCE STARTED WILL NOT TAKE MORE THAN 10 WORKING DAYS, WITH YOUR FULL SUPPORT. THIS TRANSACTIONIS 100% RISK FREE.
IF THIS PROPOSAL SATISFIES YOU, PLEASE REACH US ONLY BY FAX OR PHONE,FOR MORE INFORMATION. IT MIGHT BE DIFFICULT TO GET THROUGH TO ME, BECAUSE OF POOR TELECOMMUNICATION SYSTEM HERE. PLEASE KEEP TRYING, YOU WILL DEFINITELY GET THROUGH. PLEASE TREAT AS URGENT AND VERY CONFIDENTIAL. YOURS FAITHFULLY, DR.REGINALD AKON NB.: FOR CONFIDENTIAL REASONS AND DUE TO THE POOR COMMUNICATION SYSTEM IN MY COUNTRY, MOST OFTEN FOREIGN CALLS COULD BE DIVERTED TO THE WRONG PERSON. SO FOR YOU TO BE VERY SURE YOU ARE RIGHTLY SPEAKING WITH ME, IT IS VERY IMPORTANT THATWHEN YOU CALL AND ASK FOR ME, THE MOMENT I PICK UP THE PHONE, YOU SHOULD ASK ME FORTHE "CODE WORD" AND MY ANSWER WOULD BE "BORNGREAT" BEFORE WE PROCEED DISCUSSIONS, BUT IF I DO NOT SAY "BORNGREAT",THAT MEANS YOU ARE NOT SPEAKING WITH ME JUST DISCONNECT THE LINE AND CALL ME BACK TILL I GIVE YOU THE CODE WORD.
WE ARE MEMBERS OF A SPECIAL COMMITTEE FOR BUDGET AND PLANNING OF THE NIGERIAN NATIONAL PETROLEUM CORPORATION (NNPC). THIS COMMITTEE IS PRINCIPALLY CONCERNED WITH CONTRACT AWARDS AND APPROVAL. WITH OUR POSITIONS, WE HAVE SUCCESSFULLY SECURED FOR OURSELVES THE SUM OF TWENTY-ONE MILLION, FIVE HUNDRED THOUSAND UNITED STATES DOLLARS(US$21.5M). THIS AMOUNT WAS CAREFULLY MANIPULATED BY OVER-INVOICING OF AN OLD CONTRACT.
BASED ON INFORMATION GATHERED ABOUT YOU, WE BELIEVE YOU WOULD BE IN A POSITION TO HELP US IN TRANSFERRING THIS FUND (US$21.5M) INTO A SAFE ACCOUNT. IT HAS BEEN AGREED THAT THE OWNER OF THE ACCOUNT WILL BE COMPENSATED WITH 20% OF THE REMITTEDFUNDS, WHILE WE KEEP 70%, AND 10% WILL BE SET ASIDE TO OFFSET EXPENSES AND PAYTHE NECESSARY TAXES.
ALL MODALITIES OF THIS TRANSACTION HAVE BEEN WORKED OUT AND ONCE STARTED WILL NOT TAKE MORE THAN 10 WORKING DAYS, WITH YOUR FULL SUPPORT. THIS TRANSACTIONIS 100% RISK FREE.
IF THIS PROPOSAL SATISFIES YOU, PLEASE REACH US ONLY BY FAX OR PHONE,FOR MORE INFORMATION. IT MIGHT BE DIFFICULT TO GET THROUGH TO ME, BECAUSE OF POOR TELECOMMUNICATION SYSTEM HERE. PLEASE KEEP TRYING, YOU WILL DEFINITELY GET THROUGH. PLEASE TREAT AS URGENT AND VERY CONFIDENTIAL. YOURS FAITHFULLY, DR.REGINALD AKON NB.: FOR CONFIDENTIAL REASONS AND DUE TO THE POOR COMMUNICATION SYSTEM IN MY COUNTRY, MOST OFTEN FOREIGN CALLS COULD BE DIVERTED TO THE WRONG PERSON. SO FOR YOU TO BE VERY SURE YOU ARE RIGHTLY SPEAKING WITH ME, IT IS VERY IMPORTANT THATWHEN YOU CALL AND ASK FOR ME, THE MOMENT I PICK UP THE PHONE, YOU SHOULD ASK ME FORTHE "CODE WORD" AND MY ANSWER WOULD BE "BORNGREAT" BEFORE WE PROCEED DISCUSSIONS, BUT IF I DO NOT SAY "BORNGREAT",THAT MEANS YOU ARE NOT SPEAKING WITH ME JUST DISCONNECT THE LINE AND CALL ME BACK TILL I GIVE YOU THE CODE WORD.
On 6 Feb 2003, at 12:47, Joiner Cartwright, Jr., Ph.D. wrote:
} I run a clinical EM lab with a geriatric JEOL 100C TEMSCAN microscope. } I am campaigning for new equipment and would like to get a stand alone } TEM and SEM. .
We have three EM's (2 x TEM and 1 x SEM) in an area of about 10m x 5m, separated only by floor to ceiling curtains. This works well but remember that the air-conditioning system must be geared to handle the heat output from these instruments, peripheral PC's, etc (we have 9 monitors in that area), AND the personnel using them. We have no problem from interference (vibration or magnetic field) between the instruments.
An advantage of this arrangement is that it is very useful for exhibitions, teaching, etc.
Regards
Rob
ps. 39C here yesterday, and heading that way, or more, already today - airconditioning HAS to work well.
=====================================
Rob Cross Director : EM Unit, Rhodes University tel: (046) 603 8168/9
We have just had a look at some Poly polysulfone filters. Thanks for the list on the sample prep. We found Al, Cu, Fe, Ni, Cr particles and even a W and Bi particle as well in the unused filters! Some leftovers from the processing plant? This might be the case with your filters as well.
Mr S. H. Coetzee Electron Microscope Unit University of Botswana Private Bag 0022 Gaborone Botswana
} -----Original Message----- } From: Malis, Tom [mailto:malis-at-nrcan.gc.ca] } Sent: Friday, February 07, 2003 2:01 AM } To: 'Tindall, Randy D. '; 'microscopy-at-sparc5.microscopy.com ' } Subject: RE: ultramicrotomy: knife damage } } } -------------------------------------------------------------- } ---------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ---------. } } } Randy, I've not the foggiest as to what molecular sieves are } made of. Are } there any 'hard bits' in them, like fine ceramic particles? } If so, that } could be your problem. In many hard materials edge chipping } is a reality. } However, when sectioning uniformly hard materials like } ceramics, the only } way they will section is if one has an ultrafine facet on the } block, ie the } order of several microns. That methodology also has the } benefit of reducing } the area of damage along the edge, and one can 'walk' along } the edge for } some time before resharpening is needed. So if there is something } hard/tough enough in your sieves to chip the edge, and you are cutting } sections of hundreds of microns, you will surely get chipping } spread over } that length of the edge. The question than becomes, can your } clients live } with smaller sections? } } Tom } } -----Original Message----- } } From: Tindall, Randy D. } To: microscopy-at-sparc5.microscopy.com } Sent: 2/6/2003 10:07 AM } Subject: TEM: ultramicrotomy: knife damage } } -------------------------------------------------------------- } ---------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ---------. } } } Dear Listers, } } I'm curious as to how many people have had problems using molecular } sieves in dehydration solvents, with respect to knife damage. We seem } to be going through diamond knives at a uncomfortable rate and we're } wondering if this could be a contributing factor. We are very careful } with our knives and try to minimize contact cleaning of the edges. We } soak the knives often in the recommended solution in a commercial } cleansing unit, and the knife manufacturer has evaluated one of our } knives and confirmed that the edge is chipped, not dirty. } } As a multi-user facility, we do a LOT of ultramicrotomy on a large } variety of samples, so this may just be normal wear and tear, but we } would sure like to minimize this expense. } } Thanks much! } } Randy } } Randy Tindall } EM Specialist } Electron Microscopy Core---We're the Fun Core! } 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/ } }
Works on windows, linux, solaris, etc. Has the same fonctionnality than StarOffice 5.2, is free, and saves files in an zipped XML format. It is able to read and save in MSoffice and StarOffice formats. It have a few bugs, but not more than MS Office, and works well. It have an writer, a mathematical formules editor, drawer (lie Corel draw), a calculator like Excel, and a présentation tool like PowerPoint. I did only try the presentation soft, but a texte file has a quarter size than a Word97/2000 one, and half than a rtf.
And last but not least, it is free.
J. Faerber IPCMS-GSI (Institut de Physique et Chimie des Matériaux de Strasbourg Groupe Surface et Interfaces) 23, rue de Loess ; BP43 67034 Strasbourg CEDEX 2 France
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } I hate the way Word deals with images. It makes huge files too. I would } recommend Adobe Pagemaker or Deneba Canvas for much more controllable } combination or text and graphics. If you have Acrobat (full version) } installed you can then also export the files to pdf format and share them } easily on the web. Of course, these software packages are not so ideal for } presentations and I have not yet found an alternative to Powerpoint for } that. } } (P.S. This is not just Microsoft bashing, I wish they would do a better } job of Graphics handling in word. In my opinion, the competition is better } at this just now. As a pure Word processor I find MS Word very good). } } Best wishes } } Ian } } On Mon, 20 Jan 2003 09:30:47 -0600, } {"gary.m.brown-at-exxonmobil.com"-at-sparc5.microscopy.com} wrote: } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } } } Jeff, } } } } I, too, have had significant problems working with images imported into } } Microsoft Word. However, my software is located on the PC hard drive. The } } biggest problem that I have encountered with images in Microsoft Word } } occurs when annotating images. After the image is imported into Word, } } annotation may be done in two ways (to my knowledge): (1) Text, arrows, } } etc. may be simply superimposed over the images. The problem with this } } approach is that the annotations are not linked to the image and may not } } remain superimposed on the image if the image moves. (2) Annotations can } } also be linked (probably not the best choice of words) to the image by } } double-clicking on the image to open the image field, adding the } } annotation, then closing the image field. These annotations are permanent } } unless intentionally moved or deleted. } } } } The problems occur when one implements the second option. Comparing } } images } } before and after annotation, I found that the annotated images often } } sustained substantial changes in gray or color levels. Case in point, EDS } } maps were so badly affected that the color key was no longer correct. } } } } My solutions follow: (1) Annotate images in Adobe Photoshop before } } importing into Word. Note that the effects of lossy compression on } } annotations (blurred edges) may be pronounced. (2) Use Microsoft } } PowerPoint } } for image presentation. I have encountered no problems with image files } } in } } PowerPoint. } } } } Good luck to you in your endeavors. } } } } Cheers, } } } } "The opinions expressed are those of Gary M. Brown and do not represent } } the } } opinions of ExxonMobil Corporation nor its affiliates." } } } } Gary M. Brown } } ExxonMobil Chemical Company } } Baytown Polymers Center } } 5200 Bayway Drive } } Baytown, Texas 77520-2101 } } phone: (281) 834-2387 } } fax: (281) 834-2395 } } e-mail: Gary.M.Brown-at-ExxonMobil.com } } } } } } "Oakley, Jeff" } } {oakleyj-at-rayovac.c To: {Microscopy-at-sparc5.microscopy.com} } } om} cc: } } Subject: RE: presentation images } } 01/17/03 07:50 AM } } } } } } } } } } } } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } This same phenomenon happens with my reports in Microsoft Word. In } } addition to images darkening, they sometimes shift to other pages and/or } } change size and dimension. Adding tables and text boxes to the report } } adds } } to the fun. } } } } The software that we use is networked. Our IS department has told us } } that } } the networked software has a bug that causes these things to happen when } } file sizes increase, and that there is not a patch for it. So we have } } just } } have to deal with it. } } } } Jeff Oakley } } } } } } -----Original Message----- } } } From: Corazon D. Bucana [mailto:bucana-at-audumla.mdacc.tmc.edu] } } Sent: Thursday, January 16, 2003 11:11 AM } } To: Microscopy-at-sparc5.microscopy.com } } Subject: presentation images } } } } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } I created a Power point presentation last November 2002 consisting of } } several fluorescence micrographs. The file which was rather large ( 90 } } Mb) } } } } was left in my laptop all this time. When I opened it again this week I } } find that the images are now too dark and I needed to increase brightness } } by 3-4 clicks on the brightness icon of Power point.I increased } } brightness } } } } of all the micrographs and copied the file to a CD hoping that the image } } will not deteriorate there and then compare it with the one in my laptop } } several weeks from now. Has this happened to anyone else? Is there } } something I should have done to prevent this? } } } } Any suggestions or comments will be greatly appreciated. } } } } Cora Bucana } } } } } } } } } } } } } } } } } } } } } } -- } Ian MacLaren } Technische Universität Darmstadt } Material-und Geowissenschaften } Petersenstr. 23 } 64287 Darmstadt } Germany }
November/December 2002, Vol.10-#6, p.16: Use Adobe Acrobat to Keep Original Resolutions and to Make TIFF Files From Any Program by Jerry Sedgewick. -Matt
Matthew K. Stephenson Analytical Associate Impact Analytical 1910 West Saint Andrews Road Midland, MI 48640 (989) 832-5555 X506 stephenson-at-impactanalytical.com
-----Original Message----- } From: Kevin Frischmann [mailto:kfrisch-at-amnh.org] Sent: Thursday, February 06, 2003 5:31 PM To: Microscopy-at-sparc5.microscopy.com Cc: Dusevich, Vladimir
Sounds like you have lossy image compression enabled in Acrobat, or possibly low output resolution settings.
Lossless or uncompressed options are available in Acrobat, and should give you the high quality you need for publishing. There are quality "presets" as well; "prepress" is the high quality option if I remember correctly (or just manually reduce compression and increase output resolution).
I believe there was an article with tips on using Acrobat for scientific publishing in a recent issue of "Microscopy Today". Anyone remember what issue it was?
-Kevin ------------------------------------------------ Kevin Frischmann, Laboratory Manager Microscopy & Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
At 10:25 AM 2/6/03 -0600, "Dusevich, Vladimir" {dusevichv-at-umkc.edu} wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
November/December 2002, Vol.10-#6, p.16: Use Adobe Acrobat to Keep Original Resolutions and to Make TIFF Files From Any Program by Jerry Sedgewick. -Matt
Matthew K. Stephenson Analytical Associate Impact Analytical 1910 West Saint Andrews Road Midland, MI 48640 (989) 832-5555 X506 stephenson-at-impactanalytical.com
Sounds like you have lossy image compression enabled in Acrobat, or possibly low output resolution settings.
Lossless or uncompressed options are available in Acrobat, and should give you the high quality you need for publishing. There are quality "presets" as well; "prepress" is the high quality option if I remember correctly (or just manually reduce compression and increase output resolution).
I believe there was an article with tips on using Acrobat for scientific publishing in a recent issue of "Microscopy Today". Anyone remember what issue it was?
-Kevin ------------------------------------------------ Kevin Frischmann, Laboratory Manager Microscopy & Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
At 10:25 AM 2/6/03 -0600, "Dusevich, Vladimir" {dusevichv-at-umkc.edu} wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Randy, If you use molecular seives, you should not just dump them loose into your reagent bottles, but rather eclose them in dialysis tubing or something similar. That would eliminate particles floating free in your ethanols, etc. 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
Randy, I believe that molecular sieves contain alumina that is very detrimental to knives. The fine powder from the sieves takes a very long time to settle and is easily stirred up. Why don't you try using sodium sulfate. We have used that for years without noticeable problems. Of course, we try not to stir up the bottles or use the last 1/3 of the bottle. Rather we pour the remains together with about 1" of fresh sodium sulfate at the bottom of the bottle. Then let the bottle sit a day or two and you should be okay.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building West Lafayette, IN 47907
On 2/6/03 10:07 AM, "Tindall, Randy D." {TindallR-at-missouri.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Dear Listers, } } I'm curious as to how many people have had problems using molecular } sieves in dehydration solvents, with respect to knife damage. We seem } to be going through diamond knives at a uncomfortable rate and we're } wondering if this could be a contributing factor. We are very careful } with our knives and try to minimize contact cleaning of the edges. We } soak the knives often in the recommended solution in a commercial } cleansing unit, and the knife manufacturer has evaluated one of our } knives and confirmed that the edge is chipped, not dirty. } } As a multi-user facility, we do a LOT of ultramicrotomy on a large } variety of samples, so this may just be normal wear and tear, but we } would sure like to minimize this expense. } } Thanks much! } } Randy } } Randy Tindall } EM Specialist } Electron Microscopy Core---We're the Fun Core! } 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/ } } }
Some time ago I have bout a case of 0.5L bottles of 200 alcohol for about $2 ($2.50?) each. No more molecular sieves for me.
Vladimir
} -----Original Message----- } From: Baskin, Tobias } Sent: Thursday, February 06, 2003 1:08 PM } To: Tindall, Randy D.; microscopy-at-sparc5.microscopy.com } Subject: Re: TEM: ultramicrotomy: knife damage } } } -------------------------------------------------------------- } ---------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ---------. } } } Randy, } Some people I know keep their molecular sieve in } dialysis tubing. } } Tobias } } } ------------------------------------------------------------- } ----------- } } The Microscopy ListServer -- Sponsor: The Microscopy } Society of America } } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } ------------------------------------------------------------- } ----------. } } } } } } Dear Listers, } } } } I'm curious as to how many people have had problems using molecular } } sieves in dehydration solvents, with respect to knife } damage. We seem } } to be going through diamond knives at a uncomfortable rate and we're } } wondering if this could be a contributing factor. We are } very careful } } with our knives and try to minimize contact cleaning of the } edges. We } } soak the knives often in the recommended solution in a commercial } } cleansing unit, and the knife manufacturer has evaluated one of our } } knives and confirmed that the edge is chipped, not dirty. } } } } As a multi-user facility, we do a LOT of ultramicrotomy on a large } } variety of samples, so this may just be normal wear and tear, but we } } would sure like to minimize this expense. } } } } Thanks much! } } } } Randy } } } } Randy Tindall } } EM Specialist } } Electron Microscopy Core---We're the Fun Core! } } 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/ } } } }
I have used "insert-picture-from file" in Word and a lot of different settings for Distiller, including JPEG and ZIP compression for images, but always line scans and letter on images looked better in Word file, not in PDF.
Anyway, size of the Word 2000 file is close to the sum of image sizes (758K), and if it was up to me, I'd choose Word 2000 (not PDF) files as a standard for publication. It's too bad Microsoft do not advertise as much as Adobe its free viewers, for example this one: http://office.microsoft.com/downloads/2000/wd97vwr32.aspx
Vladimir
} } Set Distiller for printer resolution. } } Were the 4 images imported by reference or } included in the final document? Big difference. } } gary g. } } } At 08:25 AM 2/6/2003, you wrote: } } } Hi Listers, } } } } Preparing my abstract for MM'03 I have found that: } } - Two page document (4 images with superimposed EDS line scans) } } saved in Word 2000 format was 758K size. } } - Saved in Word 6.0 format (as required for uploading by } } submission instructions) it had size of 2.93M (!). } } - Saved in PDF format (Acrobat 5.0) file was just 286K, } } but line scans, still pretty visible, were looking not } } nice. } } } } I believe I should upload PDF files, since Word files } } anyway will be converted in PDF for CD publishing. But are not } } we loosing quality going digital now? } } } } 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 } } } } } } } } } -----Original Message----- } } } From: Rosemary White [mailto:rosemary.white-at-csiro.au] } } } Sent: Thursday, January 30, 2003 4:00 PM } } } To: Microscopy-at-sparc5.microscopy.com } } } Subject: Re: presentation images in Microsoft Word } } } } } } } } } -------------------------------------------------------------- } } } ---------- } } } The Microscopy ListServer -- Sponsor: The Microscopy Society } } } of America } } } To Subscribe/Unsubscribe -- Send Email to } } } ListServer-at-MSA.Microscopy.Com } } } On-Line Help } } } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } -------------------------------------------------------------- } } } ---------. } } } } } } } } } Another trick is to put the images and text into a Word } } } table, if you size } } } the cells in the table to about the size you want, the images } } } will insert } } } to fit the cell size. } } } } } } And if copying and pasting, make sure you do "Paste Special" } } } and unclick } } } the box that says "float over text". That way the images } } } always stay in } } } the same spot with respect to the surrounding text and don't } } } mysteriously } } } jump around when you insert or delete text. } } } cheers, } } } Rosemary } } } } } } } } Thanks Doug- } } } } Great tips- I'm printing them up until I memorize them. } } } } Rgds, } } } } Mike Shaw } } } } Roselle, NJ } } } } } } } } } Gary, } } } } } } } } } } MS Word is such a pain because of the way it works with } } } images. A couple } } } } } of tricks I've discovered over the years. } } } } } } } } } } 1) Place the image within a text box, rather than directly } } } on the page, it } } } } } seems to give you much greater control over the location } } } on the page. It } } } } } also makes it much easier to add text annotations that } } } stay with the image. } } } } } } } } } } } } } } } If you don't want the text box to have a border you can } } } remove it by } } } } } selecting } } } } } the box outline and look for the "paint brush" icon on the } } } Draw toolbar, } } } } } then use the down arrow and select "none". If you'd like } } } the text box to } } } } } be transparent, select the text box outline and look for } } } the "paint bucket" } } } } } icon on the Draw toolbar, then use the down arrow and } } } select "none". If } } } } } you discover its hard to find the text box border once you } } } made the edge } } } } } "invisible", first select the image with a single left } } } click (it should have } } } } } the solid black resizing "handles") and then use the } } } keyboard left or right } } } } } arrow keys and the selection with move out to the textbox } } } outline (with } } } } } black } } } } } bordered resizing "handles"). } } } } } } } } } } 2) Always use the INSERT | PICTURE | FROM FILE option as } } } opposed to copying } } } } } and pasting an image into Word. I find that the images } } } are harder to work } } } } } with if I paste them in. Also, you can insert TIFF or BMP } } } } } images into Word, } } } } } you don't have to use JPEG. } } } } } } } } } } } } } } } } } }
We operate one transmission and a couple of SEMs in one large laboratory. The individual microscopes are each enclosed in a lockable cubicle which makes it easier to control access and rermove the temptation amongst visitors to tamper. Photography and handling of film for the TEM are much easier if you have a lock on the inside - I have even occasionally used the TEM microscope cubicle as an emergency film developing darkroom (not recommended) when the main darkroom was out of commission. You could even use a red light outside linked to a safelight inside the TEM room - it is quite useful.
Our cubicles are ordinary timber painted but I suspect modern fire regulations would require specially treated materials. If the partitions are built around the microscopes then you will need some large plastic sheeting dust-covers, a lot of tape and much careful cleaning after the assembly and painting before you again expose your microscopes. Ideally have the cubicles built first, get everything cleaned a couple of times and the move the microscopes in a few days later when the dust has settled.
One thing, if you do get cubicles try to have either removable panels or very big doors (maybe even extra doors) to allow more visitors, extra equipment and most importantly access for servicing the instrument - discuss this with your service engineer if you can. Before we installed anything we got the floor specifications of each microscope, some little pictures scaled to a graph paper floor plan and could confirm that the rooms were suitable.
Good luck
Malcolm Haswell e.m. unit School of Sciences University of Sunderland UK
----- Original Message ----- } From: "Malis, Tom" {malis-at-nrcan.gc.ca}
Try putting the molecular sieve inside some dialysis tubing and seal the ends (I just used staples). You end up with a molecular sieve 'sausage' which works quite well. This is not my idea but I can't remember where I got it from. We still get damage to knives but probably from other sources because our absolute ethanol is always crystal clear.
Malcolm Haswell e.m. unit School of Sciences University of Sunderland UK
----- Original Message ----- } From: "Malis, Tom" {malis-at-nrcan.gc.ca}
On Thursday, February 6, 2003, at 07:06 AM, Lawrence Oakford wrote:
} I was wondering if anyone on the list is familiar with this stain. I } am trying to find a procedure for either making this stain or a } commercial source for the stain. It was listed in a methods section } for negative staining isolated neurofibrillary tangles (Crowther, R. } A. 1991. PNAS 88:2288) but the author did not reference a source for } the stain or a procedure. I would appreciate any help provided. } } Thanks } Dear Lawrence, I would suggest buying phosphotungstic acid and lithium hydroxide and mixing them. I have had success with other negative stains by putting 5 micro-l of the specimen and 5 micro-l of 2% stain on a grid (mixing by repeated pipetting), letting this sit for 1 min, blotting the fluid to near dryness, adding 10 micro-l of 1% stain, letting this sit for 1 min, and blotting to dryness. I would try this, but perhaps someone who has used lithium phosphotungstate has a better protocol. Yours, Bill Tivol 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
On Thursday, February 6, 2003, at 04:01 PM, Malis, Tom wrote:
} Randy, I've not the foggiest as to what molecular sieves are made of. } Are } there any 'hard bits' in them, like fine ceramic particles? } Dear Tom, Molecular sieves are made of zeolites--clays with molecule-sized cavities in their structures. I think they are aluminosilicates, but I'm no expert. Yours, Bill Tivol 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
Alternate options are Adobe Framemaker and Quark Express. These are much more industrial strength than Word. For books, dissertations, long articles, etc., these predominate over Word, IMO.
For simple documents, Word is fine. Beyond that, it fails in many ways. The down side of the other options is the steep learning curve. But once this is surmounted, Word is history. This is what has been my experience.
Since Framemaker became an Adobe product, it is tightly integrated to Acrobat. Nevertheless, any program can save or print to Distiller and have really nice results. The trick is to make the settings congruent with your desired results.
gary g.
At 06:25 AM 2/6/2003, you wrote:
} I hate the way Word deals with images. It makes huge files too. I would } recommend Adobe Pagemaker or Deneba Canvas for much more controllable } combination or text and graphics. If you have Acrobat (full version) } installed you can then also export the files to pdf format and share them } easily on the web. Of course, these software packages are not so ideal } for presentations and I have not yet found an alternative to Powerpoint } for that. } } (P.S. This is not just Microsoft bashing, I wish they would do a better } job of Graphics handling in word. In my opinion, the competition is } better at this just now. As a pure Word processor I find MS Word very good). } } Best wishes } } Ian } } On Mon, 20 Jan 2003 09:30:47 -0600, } {"gary.m.brown-at-exxonmobil.com"-at-sparc5.microscopy.com} wrote: } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Vladimir I just recently converted MS Word (2000) document with color pictures into PDF using Distiller and was not able to see the difference even the Distiller's settings was not optimal. If I understand correctly, most modern printers used "post-script" format to print. It's actually the same as PDF (yes, ADOBE's job). I mean, PDF is a sequence of command (it's not actually text or graphics), which any modern printer could recognize directly (and print). From another hand any other formats (MS Word, any pictures) should be translated into "post-script" (read PDF) to be printed. Usually it happens when you sent file to printer (printer driver did the job). So, the bottom line here: any image/document you sent to printer converted, actually into PDF (post-script). Therefore, in theory you should be able to have equal quality from PDF and let say TIFF or MS Word when printed. On the screen, they should looks differently (because PDF is actually vector graphics and TIFF for instance is bitmap). I hope it helps. Sergey
At 07:30 AM 2/7/2003, 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 Box 951737 Los Angeles, CA 90095-1737
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Thanks for all replies!
I have used "insert-picture-from file" in Word and a lot of different settings for Distiller, including JPEG and ZIP compression for images, but always line scans and letter on images looked better in Word file, not in PDF.
Anyway, size of the Word 2000 file is close to the sum of image sizes (758K), and if it was up to me, I'd choose Word 2000 (not PDF) files as a standard for publication. It's too bad Microsoft do not advertise as much as Adobe its free viewers, for example this one: http://office.microsoft.com/downloads/2000/wd97vwr32.aspx
Vladimir
} } Set Distiller for printer resolution. } } Were the 4 images imported by reference or } included in the final document? Big difference. } } gary g. } } } At 08:25 AM 2/6/2003, you wrote: } } } Hi Listers, } } } } Preparing my abstract for MM'03 I have found that: } } - Two page document (4 images with superimposed EDS line scans) } } saved in Word 2000 format was 758K size. } } - Saved in Word 6.0 format (as required for uploading by } } submission instructions) it had size of 2.93M (!). } } - Saved in PDF format (Acrobat 5.0) file was just 286K, } } but line scans, still pretty visible, were looking not } } nice. } } } } I believe I should upload PDF files, since Word files } } anyway will be converted in PDF for CD publishing. But are not } } we loosing quality going digital now? } } } } 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 } } } } } } } } } -----Original Message----- } } } From: Rosemary White [mailto:rosemary.white-at-csiro.au] } } } Sent: Thursday, January 30, 2003 4:00 PM } } } To: Microscopy-at-sparc5.microscopy.com } } } Subject: Re: presentation images in Microsoft Word } } } } } } } } } -------------------------------------------------------------- } } } ---------- } } } The Microscopy ListServer -- Sponsor: The Microscopy Society } } } of America } } } To Subscribe/Unsubscribe -- Send Email to } } } ListServer-at-MSA.Microscopy.Com } } } On-Line Help } } } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } } -------------------------------------------------------------- } } } ---------. } } } } } } } } } Another trick is to put the images and text into a Word } } } table, if you size } } } the cells in the table to about the size you want, the images } } } will insert } } } to fit the cell size. } } } } } } And if copying and pasting, make sure you do "Paste Special" } } } and unclick } } } the box that says "float over text". That way the images } } } always stay in } } } the same spot with respect to the surrounding text and don't } } } mysteriously } } } jump around when you insert or delete text. } } } cheers, } } } Rosemary } } } } } } } } Thanks Doug- } } } } Great tips- I'm printing them up until I memorize them. } } } } Rgds, } } } } Mike Shaw } } } } Roselle, NJ } } } } } } } } } Gary, } } } } } } } } } } MS Word is such a pain because of the way it works with } } } images. A couple } } } } } of tricks I've discovered over the years. } } } } } } } } } } 1) Place the image within a text box, rather than directly } } } on the page, it } } } } } seems to give you much greater control over the location } } } on the page. It } } } } } also makes it much easier to add text annotations that } } } stay with the image. } } } } } } } } } } } } } } } If you don't want the text box to have a border you can } } } remove it by } } } } } selecting } } } } } the box outline and look for the "paint brush" icon on the } } } Draw toolbar, } } } } } then use the down arrow and select "none". If you'd like } } } the text box to } } } } } be transparent, select the text box outline and look for } } } the "paint bucket" } } } } } icon on the Draw toolbar, then use the down arrow and } } } select "none". If } } } } } you discover its hard to find the text box border once you } } } made the edge } } } } } "invisible", first select the image with a single left } } } click (it should have } } } } } the solid black resizing "handles") and then use the } } } keyboard left or right } } } } } arrow keys and the selection with move out to the textbox } } } outline (with } } } } } black } } } } } bordered resizing "handles"). } } } } } } } } } } 2) Always use the INSERT | PICTURE | FROM FILE option as } } } opposed to copying } } } } } and pasting an image into Word. I find that the images } } } are harder to work } } } } } with if I paste them in. Also, you can insert TIFF or BMP } } } } } images into Word, } } } } } you don't have to use JPEG. } } } } } } } } } } } } } } } } } }
I apologise if this topic has gone off the boil but my original message was 'bounced' because my work e-mail software has a nasty habit of adding attachments if I forward a message, although not if I hit reply.
Sorry to clog up the system but I thought I should also add this. I always trim my blocks with glass knives on the ultra microtome and do a final 'facing' (smoothing) of the front of the block with a fresh area of the same or a new glass knife. I never use a diamond knife until I can achieve a smooth finish on the block with a glass knife. If I am desperate then I will either just use a glass knife or an old diamond but it's never been necessary to risk a good diamond on an unknown block this way..
Invariably the biggest problems are found at the tips of pellets where any hard crystalline deposits and broken glass tend to settle and usually deeper material is less troublesome.
Malcolm Haswell e.m. unit School of Sciences University of Sunderland UK
{snip}
} ------------------------------------------------------------------- } The Microscopy ListServer -- Sponsor: The Microscopy Society of } AmericaTo Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.ComOn-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------. } } } Dear Listers, } } I'm curious as to how many people have had problems using molecular } sieves in dehydration solvents, with respect to knife damage. We seem } to be going through diamond knives at a uncomfortable rate and we're } wondering if this could be a contributing factor. We are very careful } with our knives and try to minimize contact cleaning of the edges. } We soak the knives often in the recommended solution in a commercial } cleansing unit, and the knife manufacturer has evaluated one of our } knives and confirmed that the edge is chipped, not dirty. } } As a multi-user facility, we do a LOT of ultramicrotomy on a large } variety of samples, so this may just be normal wear and tear, but we } would sure like to minimize this expense. } } Thanks much! } } Randy } } Randy Tindall } EM Specialist } Electron Microscopy Core---We're the Fun Core! } 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/
Just thought maybe this will help someone these abstract submitting days: what I ended up doing for my M&M abstract last year, I made the 1st page (all text) in MS Word, but the 2nd page I made all in Photoshop. Photoshop now allows to add and format text. I then put the two pages together in Acrobat. Yes, I did all that after it came out bad from just inserting the picture into the Word file and distilling the whole thing. To be honest, I don't remember now what was it that I didn't like, but I remember it was unacceptable.
Vlad
-- ------------------------------------------- Vladislav V. Speransky, Ph.D. Laboratory of Structural Biology NIAMS, National Institutes of Health 50 South Drive, Room 1504 Bethesda, MD 20892-8025 Phone: 301 496-3989 Fax: 301 480-7629 E-mail: Vladislav_Speransky-at-nih.gov
Applied Optical Microscopy is just about a month away, in sunny Orlando, Florida. 3.5 days of total immersion in light microscopy, including a special focus session on digital imaging.
Remember.. this course is NOT just for chemists!
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Best regards, Barbara Foster, ACS course coordinator
Microscopy/Microscopy Education, Inc. 125 Paridon Street, Suite 102 Springfield, MA 01118 PH: 413-746-6931 FX: 413-746-9311 Web: www.MicroscopyEducation.com
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If you are using imaging software, and especially software from Zeiss like KS300 / Axiovision V06.2002, as I am using, you should take care if you use it together with the virusscanner McAfee VirusScan from Network Associates Technology (McAfee).
It seems that with the latest update version (DAT-updatefile 4246) from February 05, 2003, the program recognizes certain DLL-files required for image-capturing as infected with the virus 'HackerDefender'. These file ARE NOT infected, it's a problem of the virusscanner.
Here you will find more information about this so-called virus: http://vil.nai.com/vil/content/v_100035.htm
A part from the website from McAfee:
"Update - Feb 6th 2003: McAfee products using the 4246 DATs are incorrectly reporting certain innocent DLL files as 'trojan or variant HackerDefender'.
The innocent files affected are DLLs related to image analysis software. The following DLLs are known to be incorrectly flagged with the 4246 DATs:
MILCOR.DLL (v6.10.0.186, 626,960 bytes) MILCOR.DLL (v6.01.00.727, 590,096 bytes) MILVGA.DLL (v6.10.00.1618, 348,432 bytes) MILVGA.DLL (v6.01.00.902, 319,760 bytes) MILGEN.DLL (v6.10.00.1257, 426,256 bytes) These DLLs are completely unrelated to the HackerDefender rootkit."
Best regards and good luck,
Sven Terclavers
°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°° Sven Terclavers LM/CLSM Microscopist Center for Transgene Technology and Gene Therapy (CTG) Campus Gasthuisberg K.U.L. O&N Herestraat 49 3000 Leuven Belgium Tel. +32 16 346173 Fax. +32 16 345990 Email: Sven.Terclavers-at-med.kuleuven.ac.be °°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°
We have some problems with a PE-staining (phycoerythrin-group, B-phycoerythrin from Molecular Probes), conjugated with Cd31 to stain bloodvessels. We would like to co-stain with an FITC-conjugate, but it seems that PE is also excited by 488nm (the FITC-excitation wavelenght) instead of only by 568nm, so we cannot trust what we see because of this bleeding-through. Did any of you had the same problem and if you could solve it, how did you do it? Another problem we see is that the PE is bleached really fast. Is this normal and what can be done? Thanks in advance,
Sven Terclavers
PS. Sorry if some of you get this message double because I send it to different mailinglists.
I need retinal tissue, preferably mice. We make a low kV desktop em and some folks are belief-challenged that we image samples without staining and get excellent results.
We want to image the photoreceptors cone and rods in the various scanning modes; TEM SEM and STEM.
We are also interested in expanding our database of images from a broad range of tissue and applications.
So, it's a 2 part email;
1) does anyone know where I can get this retinal tissue ??
2) an invitation to listers who are curious about how their samples will image in our EM to contact me
Thanks,
Ephram
Ephram Shizgal Delong America info-at-delongamerica.com
one of my lab mates was using an edwards auto 306 vaccum coater and a piece of filter paper fell between the plate that seals the bell jar from the diffusion pump. we get a backing pressure fail erroe. we are unable to get into the bell jar to remove the paper. the question is does anyone know how to manulaly get into the coater. we have a service call in but would like to resolve the issue ourselves.
I have run into this in the confocal facility I run. People have brought samples that they have prepped without first consulting me and we have all sorts of strange results. PE is not a stain of choice for fluorescence microscopy. It is very popular with the flow cytometry folks where its rapid bleaching is not an issue, nor apparently is its excitation by multiple wavelengths. I would recommend almost any of the other dyes that excite around 560 in preference to PE. You will also get a stronger (enhanced) signal if you use your Cd31 as a primary Ab then tag that with a fluorescent secondary Ab, although you may have specific reasons for not wanting to do so. Try TRITC, Texas Red, Alexa 546, CY3. They will all fluoresce red and will allow your FITC double stain. 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
out of interest, as a clinical virologist who has a bit of EM background, i would be curious about the unit you are using. i have seen something marketed by one of our canadian distributors, but have no knowledge on how effective the unit might be. sorry, but i forgot the name of the manufacturer.
Does one of you know if allophycocyanin, a member of the phycobiliprotein-group (as PE), has also a high bleeching, just as PE? We have to choose a red label and can only choose between PE, APC and perCP... Thanks,
Many thanks to everyone who replied to my question about possible damage to diamond knives due to use of molecular sieves in the dehydration solvent. There doesn't seem to be a real consensus on this issue, with some folks saying they have never had a problem even after a couple of decades, and others saying they wouldn't use molecular sieves under any circumstances. The latter people advocated using freshly opened absolute ethanol (or whatever) for the final dehydration steps.
One repeated suggestion was that molecular sieves can be safely used if they are put in dialysis tubing to keep particulates safely contained. I think we'll try this.
Another suggestion made by a couple folks was that use of glass pipettes during processing could be the source of glass chips that damage knives. We never use glass pipettes in our sample preps, but interestingly enough, a client who sometimes brings us blocks prepared in his own lab does use them! Mystery solved? We'll see.
Thanks again to everyone. As always, this list is a real resource.
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We're the Fun Core! 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/
I have a small problem I hope someone can help with. We have a Lecia DMIL inverted microscope in our facility with a 6V 35W bulb for illumination. It has been using bulbs with a high frequency which surprised me as I have never changed the 100W bulbs in my Zeiss microscopes in 15 years of use. I checked the voltage output at the socket and its maximum is 10.5V. This seems odd for a 6V bulb and Leica cannot tell me if it is abnormal or not, but rates the bulbs at 50 hours which also seems odd (although that is about how long they last). It seems to me that putting a 6V bulb in a 10V socket would certainly lead to a short lifetime, but I don't know if this is our scope, bad design or something is wrong. If someone has a DMIL and can check the socket voltage, that would tell me quickly where the problem lies. Any other suggestions welcome. Thanks- Dave --
Dr. David Knecht Department of Molecular and Cell Biology University of Connecticut 91 N. Eagleville Rd. U-3125 Storrs, CT 06269-3125 knecht-at-uconn.edu 860-486-2200 860-486-4331 (fax) home page: http://www.sp.uconn.edu/~mcbstaff/knecht/knecht.html
"Electron energy loss spectroscopy (EELS) is making rapid progress in its ability to provide compositional and chemical information on the nanometre scale. A workshop on the EELS of Steels and Alloys will be held in Bruck an der Mer (Austria) from 12th - 14th June 2003 to discuss the latest developments in this area of application. The deadline for abstracts is 15th May. Details can be found at http://www.cis.tugraz.at/felmi/EELS_of_STEELS_2003.html"
Prof A J Craven Department of Physics and Astronomy University of Glasgow Glasgow G12 8QQ Scotland, UK
David: I recently went to a bulb website (Topbulb.com) looking for a bulb for my Zeiss Axiophot. They had several brands that were all the right voltage and wattage but to my amazement, the stated lifespan varied from 50 hours for some and 1000 hours for others. I think you could find a longer duration bulb with the same specs. Tom
At 11:05 AM 2/12/2003 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Associate Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
Dear Listservers, Is there anyone who has attempted to compile and use WIEN97/WIEN2k software code on an Apple Macintosh (OSX)? The code is written in FORTRAN90 and runs under Unix on 'practically all platforms'. Your comments and suggestions would be greatly appreciated.
Thanks. Mike Colella Materials & Engineering Science Australian Nuclear Science & Technology Organisation
STRICTLY CONFIDENTIAL DR. HASSAN BELLO DIRECTOR, ACCOUNTS AND AUDIT, NIGERIAN NATIONAL PETROLEUM CORPORATION (N.N.P.C) TELEOHONE NUMBER;234-8034748664 dear Sir,
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David You have to check voltage under the load, i.e. bulb should be in place and lit. If the voltage will be still high, it may be a problem. Another things just to keep in mind: does bulb has good electrical contact in the socket? It may happens even on brand new instrument (if somebody put a few drops of water into the socket - terrorist?). The sign of such problem: overheated socket, signs of oxidation/damage on the bulb's "legs" or socket. Sergey
At 08:05 AM 2/12/2003, 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 Box 951737 Los Angeles, CA 90095-1737
I have a colleague who is looking at silica particles in a polymer matrix. He is planning TEM analysis of cross sections to image the particles, determine their distribution and possibly some EDXS mapping. The samples are cast as thin films and the silica could be as small as 5nm. He has found a reference to the use of Phosphotungstic Acid (hope the spelling is correct) to stain the polymer but not the silica, thus improving contrast in the TEM. This was done by floating off the cryo-microtomed sections on a methanol solution of Phosphotungstic Acid. Alas, this is all the information he has.
We don't have cryo microtomy facilities, so we were going to use our microtome at room temp and also attempt to tripod polish.
Does anyone have a more detailed procedure of how to make up the solution and stain the polymer?? Any assistance would be greatly appreciated.
Regards George
George Theodossiou Physicist / Electron Microscopist CSIRO Manufacturing & Infrastructure Technology Private Bag 33 Clayton South MDC Victoria, 3169 tel: +61 3 9545 2012 fax: +61 3 9544 1128
To the extent permitted by law, CSIRO does not represent, warrant and/or guarantee that the integrity of this communication has been maintained or that the communication is free of errors, virus, interception or interference.
The information contained in this e-mail may be confidential or privileged. Any unauthorised use or disclosure is prohibited. If you have received this e-mail in error, please delete it immediately and notify George Theodossiou on +61 3 9545 2012. Thank you.
I am unfamiliar with your particular Lecia, but can offer some general info. Lamp life is significantly shortened under overvoltage conditions, but usually they are much bright brighter and the output spectrum moves to the shorter wavelengths. This could be intended, but not likely due to the very short life.
You need to measure the voltage under load (lamp connected and on). It could be the power supply is poorly regulated and depends on the lamp current to drop the voltage to the correct value. It this is the case, different lamps/currents would result in different loaded voltages which may or may not be correct. If the voltage does not drop, buy lamps by the case (or fix the P.S.)!
Starting incandescent lamps with an overvoltage is just the opposite of what would be the best engineering practice for life extension. Ideally (but rare), a "soft start" would be employed which limits inrush currents when the lamp is turned on. The resistance of a cold filament is very low and rises with temperature to the design value for a particular lamp. Since I=E/R (I=current, E=voltage, R=resistance), the worst time for overvoltage driven excessive current is when the filament is cold.
Another method to limit current would be a magnetically saturating transformer. I have never seen this method of current limiting used in simple lamps, but is almost always used in many welders and everyday microwave oven power supplies.
Woody White BWXT Services: http://www.bwxt.com/bwxt.html My Site: http://woody.white.home.att.net
-----Original Message----- } From: David Knecht [mailto:knecht-at-uconn.edu] Sent: Wednesday, February 12, 2003 11:06 AM To: microscopy-at-sparc5.microscopy.com
I have a small problem I hope someone can help with. We have a Lecia DMIL inverted microscope in our facility with a 6V 35W bulb for illumination. It has been using bulbs with a high frequency which surprised me as I have never changed the 100W bulbs in my Zeiss microscopes in 15 years of use. I checked the voltage output at the socket and its maximum is 10.5V. This seems odd for a 6V bulb and Leica cannot tell me if it is abnormal or not, but rates the bulbs at 50 hours which also seems odd (although that is about how long they last). It seems to me that putting a 6V bulb in a 10V socket would certainly lead to a short lifetime, but I don't know if this is our scope, bad design or something is wrong. If someone has a DMIL and can check the socket voltage, that would tell me quickly where the problem lies. Any other suggestions welcome. Thanks- Dave --
Dr. David Knecht Department of Molecular and Cell Biology University of Connecticut 91 N. Eagleville Rd. U-3125 Storrs, CT 06269-3125 knecht-at-uconn.edu 860-486-2200 860-486-4331 (fax) home page: http://www.sp.uconn.edu/~mcbstaff/knecht/knecht.html
We have had the glass in resin blocks problem recently. I only use plastic pipettes. I started suspecting the people who grew the cells but feel there has been a change in some product I use. My paranoid suspicions rest on the type of glass fracture from osmium and accelerator ampoules. Today I found an emnormous sliver 1mm long at the bottom of a block. It looks to big for molecular sieve (a suspect) EDX should indicate if it is glass. (I suppose I could do EDX on the ampoules too!).
I can cope with the problem by using a razor blade to remove the lowest part of the block.
Dave
On Wed, 12 Feb 2003 08:20:09 -0600 "Tindall, Randy D." {TindallR-at-missouri.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Many thanks to everyone who replied to my question about possible damage } to diamond knives due to use of molecular sieves in the dehydration } solvent. There doesn't seem to be a real consensus on this issue, with } some folks saying they have never had a problem even after a couple of } decades, and others saying they wouldn't use molecular sieves under any } circumstances. The latter people advocated using freshly opened } absolute ethanol (or whatever) for the final dehydration steps. } } One repeated suggestion was that molecular sieves can be safely used if } they are put in dialysis tubing to keep particulates safely contained. } I think we'll try this. } } Another suggestion made by a couple folks was that use of glass pipettes } during processing could be the source of glass chips that damage knives. } We never use glass pipettes in our sample preps, but interestingly } enough, a client who sometimes brings us blocks prepared in his own lab } does use them! Mystery solved? We'll see. } } Thanks again to everyone. As always, this list is a real resource. } } Randy } } Randy Tindall } EM Specialist } Electron Microscopy Core---We're the Fun Core! } 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/ } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
We have received from an other lab a second hand SEM with a Tracor TN 2000 unit for EDX analysis. This Tracor TN 2000 unit does not work anymore. The Tracor supplier in Europe said us that no support can be given for this product anymore. We would like to repair it by ourselves.
Can somebody help us with a copy of operator's manual and, very important for us, with the circuit diagrams? Of course, we will pay the copy and mailing costs.
Thanks a lot in advance. Best regards,
Jean Dr. Jean DILLE Materials Science and Electrochemistry Free University of Brussels, CP 194/03 Avenue F. Roosevelt, 50 1050 Brussels Belgium tel: 32-2-6502723 fax: 32-2-6502786 e-mail: jdille-at-ulb.ac.be
I seem to have missed most of the discussion about the new formulation Kodak 4489 film. Would someone please let me know what's the latest conclusion on that? I have been using the new film for several months without problem, but recently I am experiencing problems with the new film. What alternatives to Kodak film would anyone recommend?
You could try first your specimens without any staining. I have no problem in observation of silica particles and hydroxyapatite crystals (5-50 nm) embedded in adhesive resin. Particles have much greater electron density than resin and staining of the resin will only hide these particles. And for EDS its better do not apply any staining whenever possible.
Vladimir
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: "George.Theodossiou-at-csiro.au"-at-sparc5.microscopy.com } [mailto:"George.Theodossiou-at-csiro.au"-at-sparc5.microscopy.com] } Sent: Thursday, February 13, 2003 1:14 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: TEM Polymer Staining Procedure } } } -------------------------------------------------------------- } ---------- } The Microscopy ListServer -- Sponsor: The Microscopy Society } of America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -------------------------------------------------------------- } ---------. } } } Dear All, } } I have a colleague who is looking at silica particles in a } polymer matrix. } He is planning TEM analysis of cross sections to image the particles, } determine their distribution and possibly some EDXS mapping. } The samples } are cast as thin films and the silica could be as small as 5nm. } He has found a reference to the use of Phosphotungstic Acid (hope the } spelling is correct) to stain the polymer but not the silica, } thus improving } contrast in the TEM. This was done by floating off the } cryo-microtomed } sections on a methanol solution of Phosphotungstic Acid. } Alas, this is all } the information he has. } } We don't have cryo microtomy facilities, so we were going to use our } microtome at room temp and also attempt to tripod polish. } } Does anyone have a more detailed procedure of how to make up } the solution } and stain the polymer?? Any assistance would be greatly } appreciated. } } } Regards } George } } George Theodossiou } Physicist / Electron Microscopist } CSIRO Manufacturing & Infrastructure Technology } Private Bag 33 Clayton South MDC } Victoria, 3169 } tel: +61 3 9545 2012 } fax: +61 3 9544 1128 } } Visit our Web site http://www.cmst.csiro.au } } Shipping address: CSIRO - Manufacturing & Infrastructure } Technology, Gate 4 } Normanby Rd. Clayton, Victoria, } } PLEASE NOTE: } } To the extent permitted by law, CSIRO does not represent, } warrant and/or } guarantee that the integrity of this communication has been } maintained or } that the communication is free of errors, virus, interception or } interference. } } The information contained in this e-mail may be confidential } or privileged. } Any unauthorised use or disclosure is prohibited. If you } have received this } e-mail in error, please delete it immediately and notify } George Theodossiou } on +61 3 9545 2012. Thank you. } } } }
Picking up Woody's point of 'slow-starting' microscope bulbs, a simple way to achieve this with DC bulbs is to connect a hefty capacitor in parallel with the bulb. This gives a gradual switching on and off of the bulb, the speed depending upon the size of the capacitor. We employ this technique for all our small, undergraduate benchtop microscopes as they tend to get switched on and off a lot. This might not be the solution to the problem under discussion but it's very quick, easy and cheap to try and straightforward to undo if it doesn't do the trick.
Justin
} Starting incandescent lamps with an overvoltage is just the opposite of what } would be the best engineering practice for life extension. Ideally (but } rare), a "soft start" would be employed which limits inrush currents when } the lamp is turned on. The resistance of a cold filament is very low and } rises with temperature to the design value for a particular lamp. Since } I=E/R (I=current, E=voltage, R=resistance), the worst time for overvoltage } driven excessive current is when the filament is cold.
---------------------------------- Justin Ritherdon, Materials Science and Engineering, Department of Engineering, University of Liverpool, LIVERPOOL, L69 3GH, United Kingdom
We have had a somewhat similar problem with different make/models of scopes and were told by the scope repair facility that transformers and particularly rheostats anywhere in the electrical system will age and become unreliable. This unreliability results in "spikes" and variations in voltage/power to the bulb and greatly shortens their life. Their suggestion was to rebuild any rheostat when bulb life becomes a issue. On at least one scope with a consistent history of burn out, he was able to demonstrate to us the very brief flickering that the bulb exhibited as it was first turned on and stepped up through the variable power ranges.
bye for now
Tom Parker CSDLAC {tparker-at-lacsd.org}
-----Original Message----- } From: David Knecht [SMTP:knecht-at-uconn.edu] Sent: Wednesday, February 12, 2003 8:06 AM To: microscopy-at-sparc5.microscopy.com
I have a small problem I hope someone can help with. We have a Lecia DMIL inverted microscope in our facility with a 6V 35W bulb for illumination. It has been using bulbs with a high frequency which surprised me as I have never changed the 100W bulbs in my Zeiss microscopes in 15 years of use. I checked the voltage output at the socket and its maximum is 10.5V. This seems odd for a 6V bulb and Leica cannot tell me if it is abnormal or not, but rates the bulbs at 50 hours which also seems odd (although that is about how long they last). It seems to me that putting a 6V bulb in a 10V socket would certainly lead to a short lifetime, but I don't know if this is our scope, bad design or something is wrong. If someone has a DMIL and can check the socket voltage, that would tell me quickly where the problem lies. Any other suggestions welcome. Thanks- Dave --
Dr. David Knecht Department of Molecular and Cell Biology University of Connecticut 91 N. Eagleville Rd. U-3125 Storrs, CT 06269-3125 knecht-at-uconn.edu 860-486-2200 860-486-4331 (fax) home page: http://www.sp.uconn.edu/~mcbstaff/knecht/knecht.html
Another good bulb website: bulbman.com. John Mardinly Intel
-----Original Message----- } From: Tom Phillips [mailto:phillipst-at-missouri.edu] Sent: Wednesday, February 12, 2003 10:11 AM To: David Knecht Cc: Microscopy-at-sparc5.microscopy.com
David: I recently went to a bulb website (Topbulb.com) looking for a bulb for my Zeiss Axiophot. They had several brands that were all the right voltage and wattage but to my amazement, the stated lifespan varied from 50 hours for some and 1000 hours for others. I think you could find a longer duration bulb with the same specs. Tom
At 11:05 AM 2/12/2003 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thomas E. Phillips, PhD Associate Professor of Biological Sciences Director, Molecular Cytology Core 3 Tucker Hall University of Missouri Columbia, MO 65211-7400
Just to give an update. I had been experiencing so called fogging problem with the new formulation of the kodak 4489 film.
Somehow, I can't pinpoint why but my problem has disappeared. I am still using the new formulation film.
Kodak tells me that I was perhaps not agitating enough or properly. I've developed some film without any agitation and I still don't get any fogging (not that I want it back).
Maybe I had a batch before who knows.
Rajesh Patel Robert Wood Johnson Medical School Department of Pathology 675 Hoes Lane Piscataway, NJ 08854
On our 306A there is a bleed valve meant for plasma-glo. When the diffusion pump is cold, open the bleed valve and wait. I think it took about 2hrs to come to atmosphere. Then you can open the bell jar and remove the diffusion pump valve(unscrew like a piano stool). Hopefully your filter paper will be reachable from there.
Ed Hurley
Roswell Park Cancer Institute Corporation
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We also have been struggling with this new formulation for 3 weeks now. The emulsion setting is apparently unchanged [sensitivity to photons], but the film requires much, much more agitation [both more frequent motion, and larger motions help] to get an approximately even development. It is as if the film clings to the developer in a patchy fashion, so that one tends to get very muddy, uneven images. It is almost impossible to get good development near the edge of the film holder, apparently due to poor exchange of solutions, even with lots of agitation. We are also being extra careful to fully drain all developer from the film before going to the 1st rinse, to agitate a little during this rinse, and to agitate a lot in the fix step, again using larger motions, and more frequently.
It is really easy with this film to obtain terrible, unprintable images. But with more concentrated efforts by the user, most of the problem can be dealt with. We've wasted at least one hundred shots getting our methods corrected, and produced some really ugly images for users during January. Hope to be back on track soon. Would love to know how to get an even development all the way to the edge of the film.
We would be interested to hear if people using a nitrogen burst system find any problems? Is that sufficient to deal with the changes in film properties? -- David H. Hall, Ph.D. Center for C. elegans Anatomy Department of Neuroscience Albert Einstein College of Medicine 1410 Pelham Parkway Bronx, NY 10461
TO ALL: WE ARE TRYING TO OBTAIN SOME LAB-6 PARTS FOR AN ISI-WB6, SO IF YOU ARE DECOMMISIONING OR SCRAPPING ONE OF THESE MODEL "SEM'S". PLEASE CONTACT US OFF LINE.
Dear Colleagues, The 4th ASEAN Microscopy Conference (ASEANMC4) will be held in Hanoi, VietNam, from October 9 till 10, 2003. This Conference will be a gathering of microscopists from the ASEAN and other regions of the world. An important aim of the Conference is to promote mutual understandings and to develop international collaborative research in life and material sciences. Participants will have the opportunity to review recent work, to discuss recent advances and identify new challenges in the field of electron and light microscopy. The Second announcement are currently in the mail to many microscopist who are interested in this conference. When we receive reply from you then we will send "the Second announcement" to you by post as soon as possible. We would like to extend our warmest welcome to you to participate and look forward to seeing you in Hanoi in October 2003. Chairman, Organizing committee Prof. Nguyen Van Man
*********************************************** Mailing address: Assoc. Prof. Nguyen Kim Giao Electron Microscopy Unit National Institute of Hygiene and Epidemiology N0 1, Yersin street- Hai Ba Trung district - Hanoi - Vietnam Tel: 84.04.9715434 Fax: 84.04.8210853 Email: emlad-at-hn.vnn.vn, or emunihe-at-vol.vnn.vn ***********************************************
Institute of Physics EMAG 2003 Electron Microscopy and Analysis Group Conference 2003 "Towards Objective Oriented Microscopy" The University of Oxford, UK, 3 - 5 September 2003 Call for Papers
Dates for your Diary 21 March 2003 Deadline for Abstract Submission April 2003 Abstract Notification May 2003 Registration Mailing 18 July 2003 Deadline for Registration
General Enquiries: WWW: http://physics.iop.org/IOP/Confs/EMG/ For further information please contact Jasmina Bolfek- Radovani,The Institute of Physics, 76 Portland Place, London W1B 1NT, Tel: +44 (0)20 7470 4800 Fax: +44 (0)20 7470 4900, Email: jasmina.bolfek-radovani-at-iop.org
Exhibition Enquiries: Ron Doole, EMAG03, Department of Materials, University of Oxford, Parks Road, Oxford., OX1 3PH Tel: + 44 (0)1865 273701 Fax: + 44 (0)1865 283333 Email:ron.doole-at-materials.oxford.ac.uk
We're using a nitrogen burst system and haven't had any problems with our 4489 films. Actually I was wondering what all the fuss was about! Maybe that's the solution. Good luck
Marc
On Thursday, February 13, 2003, at 04:33 PM, David Hall wrote:
} ----------------------------------------------------------------------- } - } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ----------------------------------------------------------------------- } . } } } We also have been struggling with this new formulation for 3 weeks } now. The emulsion setting is apparently unchanged [sensitivity to } photons], but the film requires much, much more agitation [both more } frequent motion, and larger motions help] to get an approximately even } development. It is as if the film clings to the developer in a patchy } fashion, so that one tends to get very muddy, uneven images. It is } almost impossible to get good development near the edge of the film } holder, apparently due to poor exchange of solutions, even with lots } of agitation. We are also being extra careful to fully drain all } developer from the film before going to the 1st rinse, to agitate a } little during this rinse, and to agitate a lot in the fix step, again } using larger motions, and more frequently. } } It is really easy with this film to obtain terrible, unprintable } images. But with more concentrated efforts by the user, most of the } problem can be dealt with. We've wasted at least one hundred shots } getting our methods corrected, and produced some really ugly images } for users during January. Hope to be back on track soon. Would love } to know how to get an even development all the way to the edge of the } film. } } We would be interested to hear if people using a nitrogen burst system } find any problems? Is that sufficient to deal with the changes in } film properties? } -- } David H. Hall, Ph.D. } Center for C. elegans Anatomy } Department of Neuroscience } Albert Einstein College of Medicine } 1410 Pelham Parkway } Bronx, NY 10461 } } www.wormatlas.org } www.aecom.yu.edu/wormem } } phone 718 430-2195 } fax 718 430-8821 } }
-- Marc Pypaert Department of Cell Biology Center for Cell and Molecular Imaging Ludwig Institute for Cancer Research Yale University School of Medicine 333 Cedar Street, PO Box 208002 New Haven, CT 06520-8002 TEL 203-785 3681 FAX 203-785 7446
I am a hydrophobe when it comes to Araldite and Spurrs. I was taught that sticky blocks were often due to water in the ethanol. I have never tried experiments to check this advice.
Dave
On Fri, 14 Feb 2003 10:26:55 -0500 Geoff McAuliffe {mcauliff-at-UMDNJ.EDU} wrote:
} On the subject of molecular seives and keeping our absolute ethanol } water-free I was always taught that every last molecule of water had to be } removed during processing for EM. Well, it turns out that Epon is miscible } with 70% ethanol! Check out Hayat's Principles and Techniques of EM, } biological applications, vol. 1, page 154. Also, one of my colleagues } routinely goes from 95% ethanol to Epon (ok, an Epon substitute, I don't know } which one) with no problems. I don't know about Araldite, Spurr, etc. } } Geoff } } } On Wed, 12 Feb 2003 08:20:09 -0600 "Tindall, Randy D." } } {TindallR-at-missouri.edu} wrote: } } } } } Many thanks to everyone who replied to my question about possible damage } } } to diamond knives due to use of molecular sieves in the dehydration } } } solvent. There doesn't seem to be a real consensus on this issue, with } } } some folks saying they have never had a problem even after a couple of } } } decades, and others saying they wouldn't use molecular sieves under any } } } circumstances. The latter people advocated using freshly opened } } } absolute ethanol (or whatever) for the final dehydration steps. } } } } } } One repeated suggestion was that molecular sieves can be safely used if } } } they are put in dialysis tubing to keep particulates safely contained. } } } I think we'll try this. } } } } } } Another suggestion made by a couple folks was that use of glass pipettes } } } during processing could be the source of glass chips that damage knives. } } } We never use glass pipettes in our sample preps, but interestingly } } } enough, a client who sometimes brings us blocks prepared in his own lab } } } does use them! Mystery solved? We'll see. } } } } } } Thanks again to everyone. As always, this list is a real resource. } } } } } } Randy } } } } } } Randy Tindall } } } EM Specialist } } } Electron Microscopy Core---We're the Fun Core! } } } 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/ } } } } } -- } ********************************************** } 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 } ********************************************** }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
I have a class lecture to give next week on localization, but I don't have an image (nor do I seem to be able to readily locate one) of a GFP molecular model. I was hoping some one out there would have one I could use (digital just good enough for electronic power point) or point me in the right direction.
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
There has been a great deal of discussion about the problems with the newly formatted Kodak type 4489 film. We have been using the Kodak type S-063 for many years without incident. It is more similar to the old Electron Image plates that we used through the late 80¹s. Now I understand that S-063 is faster than 4489 and has slightly larger grain but it also can stand more manipulation regarding exposure and development times. Small film grain is important. However, unless you are doing very critical high magnification imaging, I wonder if the slightly larger grain in S-063 would be a problem.
Has anyone done a comparison between the two films recently? This may be an alternative that you can try.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building West Lafayette, IN 47907 On 2/13/03 9:03 AM, "Eleana Sphicas" {sphicae-at-rockefeller.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } I seem to have missed most of the discussion about the new formulation } Kodak 4489 film. Would someone please let me know what's the latest } conclusion on that? I have been using the new film for several months } without problem, but recently I am experiencing problems with the new film. } What alternatives to Kodak film would anyone recommend? } } Eleana } } }
Hi All: I have been asked by a colleague to ask everyone two things: 1) Is there anyone with a Philips 430 that they are parting out?
2) Is there anyone that has been able to align the Philips 430 without the use of the alignment controls (many of his microscope's electronic alignment controls are non operational). If so, please contact me off-line with ideas/insights or techniques. Thanks in advance, Michael Coviello UT Arlington
Since I haven't had the problem YET, but likely will soon, IF I do see what you have seen, I'll likely try a little PhotoFlo in the developer to help with film wettability.
Fred Monson
-----Original Message----- } From: David Hall [mailto:hall-at-aecom.yu.edu] Sent: Thursday, February 13, 2003 4:33 PM To: Microscopy-at-sparc5.microscopy.com
We also have been struggling with this new formulation for 3 weeks now. The emulsion setting is apparently unchanged [sensitivity to photons], but the film requires much, much more agitation [both more frequent motion, and larger motions help] to get an approximately even development. It is as if the film clings to the developer in a patchy fashion, so that one tends to get very muddy, uneven images. It is almost impossible to get good development near the edge of the film holder, apparently due to poor exchange of solutions, even with lots of agitation. We are also being extra careful to fully drain all developer from the film before going to the 1st rinse, to agitate a little during this rinse, and to agitate a lot in the fix step, again using larger motions, and more frequently.
It is really easy with this film to obtain terrible, unprintable images. But with more concentrated efforts by the user, most of the problem can be dealt with. We've wasted at least one hundred shots getting our methods corrected, and produced some really ugly images for users during January. Hope to be back on track soon. Would love to know how to get an even development all the way to the edge of the film.
We would be interested to hear if people using a nitrogen burst system find any problems? Is that sufficient to deal with the changes in film properties? -- David H. Hall, Ph.D. Center for C. elegans Anatomy Department of Neuroscience Albert Einstein College of Medicine 1410 Pelham Parkway Bronx, NY 10461
We have had several reports today that nitrogen burst systems are sufficient to overcome the film's poor wettability, using 2 sec bursts every 8 seconds.
One lab has reported that a pre-rinse in water before the developer also helps to give a more even effect.
We have not had a chance yet to try these ideas [and we don't own a nitrogen burst system], but the problem is serious for anyone lacking nitrogen burst. Clearly the physics at the film surface are quite different to what we are used to.
DHH
} Since I haven't had the problem YET, but likely will soon, IF I do see what } you have seen, I'll likely try a little PhotoFlo in the developer to help } with film wettability. } } Fred Monson
-- David H. Hall, Ph.D. Center for C. elegans Anatomy Department of Neuroscience Albert Einstein College of Medicine 1410 Pelham Parkway Bronx, NY 10461
My response to David Hall's concerns about film agitation during dewvelopment:
Years ago another EM Lab and mine explored this issue. We did all kinds of agitations and came to the conclusion that very minimal agitation produced VERY even development. We exposed Kodak neg emulsions for testing under a darkroom enlarger set up to provide very even illumination and a medium developed density, so did not test using TEM exposed negs of a stained section,for example.
Minimum agitation consisted of the following procedure: at beginning, lower rack of negs into developer (we use full strength Kodak D-19), immediately raise completely out of developer and lower, 2x times. Wait 1 minute, raise out of developer 1x, lower again, wait 1 minute and repeat 1x. Wait 30 sec, raise out, lower. Wait 30 sec, remove and drain 15 sec, place into stop bath. Total time is 3.0 minutes, plus 15 sec drain time.
The 3.0 min development time was empirically determined by running exposure series on TEM (different camera speeds), photographing typical stained biological section (what we look at 90% of time).
Other minimal agitation methods would probably work too, the point is to do just 1 up and down about every minute. Of course the point of agitation is to bring fresh developer to the surface of the emulsion and to remove reaction products from that surface. Perhaps this minimal agitation method means you need a bit more time to overcome slowed development rate due to reaction product buildup between agitations, but one up/down agitation seems sufficient to replenish fresh developer at emulsion surface soon enough to give good development in 3 minutes, full strength D-19, room temp (~22C).
We concluded that the rapid and frequent agitations produce whorls of quick moving developer over the central areas of the emulsions, but create areas near the edges of the film where there is relatively reduced motion of developer and reduced development rates, due to fluid friction in the narrow spacing of about 1/8 inch between films in the rack, developer "boxed in" to the corners between tank sides, edges of films, etc. We would see reduced edge development using those agitations, uneven development across central areas of fillms. For comnparison, we did tray development, few films at a time, with agitation done by moving fingers over the films under development, also got even development that way, as no edge/corner restriction of developer flow in a tray, but who would ever develop 56 filams in a tray! Got to get it to work in a rack in a tank.
Never tried nitrogen burst system, may be fine if you have huge amounts of film to be developed daily, but seems like unnessesary compliction and expense here, given amounts of film we process weekly.
-- Gib Ahlstrand, Scientist Electron Optical Facility, University of Minnesota, CBS Imaging Center, 35 Snyder Hall, St. Paul, MN. USA. 55108 (612)624-3454 (612)625-5754 FAX, ahlst007-at-tc.umn.edu http://www.cbs.umn.edu/ic/
"You can learn a lot by observation - just by lookin'!" - Yogi Bera
} We also have been struggling with this new formulation for 3 weeks } now. The emulsion setting is apparently unchanged [sensitivity to } photons], but the film requires much, much more agitation [both more } frequent motion, and larger motions help] to get an approximately } even development. It is as if the film clings to the developer in a } patchy fashion, so that one tends to get very muddy, uneven images. } It is almost impossible to get good development near the edge of the } film holder, apparently due to poor exchange of solutions, even with } lots of agitation. We are also being extra careful to fully drain } all developer from the film before going to the 1st rinse, to agitate } a little during this rinse, and to agitate a lot in the fix step, } again using larger motions, and more frequently. } } It is really easy with this film to obtain terrible, unprintable } images. But with more concentrated efforts by the user, most of the } problem can be dealt with. We've wasted at least one hundred shots } getting our methods corrected, and produced some really ugly images } for users during January. Hope to be back on track soon. Would love } to know how to get an even development all the way to the edge of the } film. } } We would be interested to hear if people using a nitrogen burst } system find any problems? Is that sufficient to deal with the } changes in film properties?
} David H. Hall, Ph.D. } Center for C. elegans Anatomy } Department of Neuroscience } Albert Einstein College of Medicine } 1410 Pelham Parkway } Bronx, NY 10461
Does anyone have a clever easy way to separate new glass slides when you open the box and they are perfectly stuck together. I have heated them in water but that didn't work. I've tapped them on counters, picked at their corners with forceps and fingers....the glass is so clean that they are just "welded" together along their entire surface.
: "You can learn a lot by observation - just by lookin'!" - Yogi Bera : : } We also have been struggling with this new formulation for 3 weeks : } now. The emulsion setting is apparently unchanged [sensitivity to : } photons], but the film requires much, much more agitation [both more : } frequent motion, and larger motions help] to get an approximately : } even development. It is as if the film clings to the developer in a : } patchy fashion, so that one tends to get very muddy, uneven images. : } It is almost impossible to get good development near the edge of the : } film holder, apparently due to poor exchange of solutions, even with : } lots of agitation. We are also being extra careful to fully drain : } all developer from the film before going to the 1st rinse, to agitate : } a little during this rinse, and to agitate a lot in the fix step, : } again using larger motions, and more frequently. : } : } It is really easy with this film to obtain terrible, unprintable : } images. But with more concentrated efforts by the user, most of the : } problem can be dealt with. We've wasted at least one hundred shots : } getting our methods corrected, and produced some really ugly images : } for users during January. Hope to be back on track soon. Would love : } to know how to get an even development all the way to the edge of the : } film. : } : } We would be interested to hear if people using a nitrogen burst : } system find any problems? Is that sufficient to deal with the : } changes in film properties? : : } David H. Hall, Ph.D.
In 40 years of developing film the most consistently even developing of sheet film I ever got was using tubes partially filled with developer constantly agatiteted while floating in water. The commercial version are BZST film tubes http://viewcamerastore.com/catalog/default.php?cPath=27&PHPSESSID=daba0c60b4 7905b73d4ee127ee91c818. I made my own tubes from black sewer pipe.
The most you can do at one time are six negatives and it requires a person full time during the development. You only need to be in the dark while loading the tubes and filling with developer.
I don't know how 4489 would respond to constant agitation but most films can be handled by lowering the developing time by up to a third and or lowering the concentration of the developer. Be careful with dilute developers because you can have more film than you have developer and exhaust the developer before the film is developed.
If you are only doing a limited number of negatives it might be worth looking into. It takes up little space and no dedicated space. Compared to a nitrogen agitations system the price is very low.I would not want to spend 4 hours a day at this. But for one or two negatives it is really great and not bad for a couple of dozen. In a pinch you can load the film and developer in a dark bag. Being completely portable. If you do go probable take you own chemicals. If you don't you will regret it some day.
Expect to do some experiments wiht development.
Good luck
Gordon Couger gcouger-at-couger.com
I collect links on information related to light microscopes. http://www.couger.com/microscope/links/gclinks.html Please forward any links or information you think might be useful to others.
No, I don't know a way. Suggest that you contact the manufacturer for any tips. Then, please report them back to us.
Regards,
"The statements and opinions expressed here by Gary M. Brown represent neither those of ExxonMobil Corporation nor its affiliates."
Gary M. Brown ExxonMobil Chemical Company Baytown Polymers Center 5200 Bayway Drive Baytown, Texas 77520-2101 phone: (281) 834-2387 fax: (281) 834-2395 e-mail: Gary.M.Brown-at-ExxonMobil.com
Tom Parker {tparker-at-lacsd.org To: "microscopy-at-sparc5.microscopy.com" } {microscopy-at-sparc5.microscopy.com} cc: Subject: glass slides 02/14/03 05:54 PM Please respond to "tparker-at-lacsd.org "
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Greetings Microscopers:
Does anyone have a clever easy way to separate new glass slides when you open the box and they are perfectly stuck together. I have heated them in water but that didn't work. I've tapped them on counters, picked at their corners with forceps and fingers....the glass is so clean that they are just "welded" together along their entire surface.
Hello List } } } } I'd like to hear Your opinion/experience about cleaning of the diamond } } indentor like Vickers or Berkovich type used in nano and micro indentation } } instruments. } } They get contaminated mainly by metals or surface deposits. Ultrsonic } } tratment can cause breaks of it or loose the mounting to holder. } } I was thinking about some plasma cleaning method but with some reactive } } media ?? what would be adviced to diamond (holded in steel mount). } } Maybe some places offer repolishing ?? } } } } regards } } } } Krzysztof Herman } } ================================= } } LABSOFT, PL 02-892 Warszawa, ul.Bazancia 45A } } =================================
At 03:31 AM 2/15/2003, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-----Original Message----- } From: Michael O'Keefe [mailto:MAOKeefe-at-lbl.gov] Sent: Thursday, February 06, 2003 7:52 PM To: Anderson, Ron
Sounds like you have lossy image compression enabled in Acrobat, or possibly low output resolution settings.
Lossless or uncompressed options are available in Acrobat, and should give you the high quality you need for publishing. There are quality "presets" as well; "prepress" is the high quality option if I remember correctly (or just manually reduce compression and increase output resolution).
I believe there was an article with tips on using Acrobat for scientific publishing in a recent issue of "Microscopy Today". Anyone remember what issue it was?
-Kevin ------------------------------------------------ Kevin Frischmann, Laboratory Manager Microscopy & Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
At 10:25 AM 2/6/03 -0600, "Dusevich, Vladimir" {dusevichv-at-umkc.edu} wrote: } ----------------------------------------------------------------------- - } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
We use a Knoop hardness tester and check the size of the dent on an SEM. This may sound simple, but scotch tape was the way we cleaned very fine diamond heads at RCA laboratories. Roy Nelson Material Testing Laboratory mtl-at-njcc.com
"Krzysztof M.Herman" wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Hello List } } } } } } I'd like to hear Your opinion/experience about cleaning of the diamond } } } indentor like Vickers or Berkovich type used in nano and micro } indentation } } } instruments. } } } They get contaminated mainly by metals or surface deposits. Ultrsonic } } } tratment can cause breaks of it or loose the mounting to holder. } } } I was thinking about some plasma cleaning method but with some reactive } } } media ?? what would be adviced to diamond (holded in steel mount). } } } Maybe some places offer repolishing ?? } } } } } } regards } } } } } } Krzysztof Herman } } } ================================= } } } LABSOFT, PL 02-892 Warszawa, ul.Bazancia 45A } } } =================================
I've had knife damage from molecular sieves, from glass chips from glass pipets, from cultured cells sscraped off coverslips, and from tissues that had been broken up with ground glass homogenizers. Most of these have been from blocks that were prepared by clients who did not check their protocol with me first!
As do others, we use fairly freshly opened pint bottles of absoute ethanol. We guesstimate when they are no longer close enough to absolute to be useful and dowgrade them to "about 95%". Life's been much easier!
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
Back in the days when we used to clean glass slides with 99% ethanol we were also able to solve the problem of slide clumps. Just leave them in the alcohol for a while and they should/will slide apart.
Let me know how you get on??
All the best.
Gareth
At 15:54 2003-02-14 -0800, Tom Parker wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Obs/NB New postal/visiting address from July 2002!
Gareth Morgan MPhil MSc FIBMS, Department of Laboratory Medicine (Labmed), Karolinska Institutet, Huddinge Universitetssjukhus, F46 SE 141 86 Stockholm Sweden
OBS! Besöksadress: F-Huset, Forskningsgatan 2 F52, Rum 2.10. Laboratoriet för klinisk patologi och cytologi.
NB! Visiting address: Building F, Research Corridor 2 F52, Room 2.10. Clinical Histo- and Cytopathology Laboratory.
I use the edge of a straight edge razor blade. This seperates them and you don't have to worry about paper towel residue.
} } } Gareth Morgan {Gareth.Morgan-at-impi.ki.se} 02/17/03 03:52AM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi
Back in the days when we used to clean glass slides with 99% ethanol we
were also able to solve the problem of slide clumps. Just leave them in the alcohol for a while and they should/will slide apart.
Let me know how you get on??
All the best.
Gareth
At 15:54 2003-02-14 -0800, Tom Parker wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} water but } that didn't work. I've tapped them on counters, picked at their corners with } forceps and fingers....the glass is so clean that they are just "welded" } together along their entire surface. } } Thanks for any suggestions. } } Tom Parker } {tparker-at-lacsd.org}
Obs/NB New postal/visiting address from July 2002!
Gareth Morgan MPhil MSc FIBMS, Department of Laboratory Medicine (Labmed), Karolinska Institutet, Huddinge Universitetssjukhus, F46 SE 141 86 Stockholm Sweden
OBS! Besöksadress: F-Huset, Forskningsgatan 2 F52, Rum 2.10. Laboratoriet för klinisk patologi och cytologi.
NB! Visiting address: Building F, Research Corridor 2 F52, Room 2.10. Clinical Histo- and Cytopathology Laboratory.
Dear Tom, I don't know how to separate glass slides stuck together, but the slides we get are "interleaved", which means there is a piece of fine paper between the slides. Mary ----- Original Message ----- } From: "Tom Parker" {tparker-at-lacsd.org} To: {microscopy-at-sparc5.microscopy.com} Sent: Friday, February 14, 2003 3:54 PM
Tom Glass cutters use a light oil to keep the split open. Obviously oil would contaminate slides, but you could try immersion in a volatile fluid, such as petroleum spirit, xylene or ethanol. Chris
Dr. Chris Jeffree Inveresk Cottage 26, Carberry Road Inveresk Musselburgh Midlothian EH21 8PR Tel: +44 131 665 6062 FAX +44 131 653 6248 Mobile 07710 585 401 ----- Original Message ----- } From: "Tom Parker" {tparker-at-lacsd.org} To: {microscopy-at-sparc5.microscopy.com} Sent: Friday, February 14, 2003 11:54 PM
I just put the on a hotplate for a second and that usually separates them... MC
Mary Mager wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Dear Tom, } I don't know how to separate glass slides stuck together, but the slides we } get are "interleaved", which means there is a piece of fine paper between } the slides. } Mary } ----- Original Message ----- } } } From: "Tom Parker" {tparker-at-lacsd.org} } } To: {microscopy-at-sparc5.microscopy.com} } Sent: Friday, February 14, 2003 3:54 PM } Subject: glass slides } } } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I need the adapter from a Link detector (c. 1990) to get my good Link detector onto a recently acquired 820 SEM. Has anyone got a dead Link detector from a JEOL 820 SEM. Even one from a good detector that's not mounted would be useful if I can borrow it long enough for my machinist to can make it.
Owen
Owen P. Mills Electron Optics Engineer Materials Science & Engineering Michigan Technological University Rm 512 M&M Bldg. Houghton, MI 49931 PH 906-369-1875 FAX 906-487-2934 mailto:opmills-at-mtu.edu http://www.mm.mtu.edu/~opmills
A method I have used to separate glass slides that are stuck together is to dip them in liquid nitrogen. They snap apart easily. This is used routinely in the histology lab when pathologists have stacked the slides while still wet with mounting media and the mounting media glues them together. I have not tried this for your application but it won't hurt to try it.
Cheryl Rehfeld Meyer Instruments, Inc. Houston, TX ----- Original Message ----- } From: Tom Parker {tparker-at-lacsd.org} To: {microscopy-at-sparc5.microscopy.com} Sent: Friday, February 14, 2003 5:54 PM
Hi all,
We have some OC darkroom filters which are going bad (cracked layers) and I would like to replace them. Unfortunately, our fixtures are 9"x9" and all the filters I've found thus far have been 8x10 or 10x12. Before I get the larger ones and cut them down, is anyone aware of a vendor who sells the 9x9 filters?
TIA, Henk Colijn
Hendrik O. Colijn colijn.1-at-osu.edu Campus Electron Optics Facility Ohio State University (614) 292-0674 http://www.ceof.ohio-state.edu Time is that quality of nature which keeps events from happening all at once. Lately it doesn't seem to be working.
Try placing slides in warm oven. Perhaps slight expansion will help separate the slides. I haven't tried this approach, so I don't know whether it will work or not.
Gary Gill
-----Original Message----- } From: Tom Parker [mailto:tparker-at-lacsd.org] Sent: Friday, February 14, 2003 6:55 PM To: microscopy-at-sparc5.microscopy.com
Greetings Microscopers:
Does anyone have a clever easy way to separate new glass slides when you open the box and they are perfectly stuck together. I have heated them in water but that didn't work. I've tapped them on counters, picked at their corners with forceps and fingers....the glass is so clean that they are just "welded" together along their entire surface.
This is to inform you that the Call for Abstracts for the forthcoming International Meeting on Applied Physics (APHYS-2003), to be held during October 14-18th 2003 in Badajoz (Spain), is now opened. All the information regarding this interdisciplinary conference can be found at the Conference website
In addition to the regular Scientific Program, several International Workshops will be held as pre-conference events. The following three Workshops are presently confirmed:
1. Workshop on Modern Applied Microscopy in Molecular and Cell Biophysics Research 2. International Interdisciplinar Workshop on Bioengineered Non-crystalline Solids 3. International Workshop on Occupational Radiation Protection
The Conference will be specifically interested in receiving reports on Interdisciplinary researches relating Physics with other Sciences such as Biology, Chemistry, Information Technology, Medicine, etc or relating different Physics areas. In other words, we are specially (but not exclusivelly) interested in reports applying the techniques, the training, and the culture of physics to research areas usually associated with other scientific and engineering disciplines
APHYS-2003 will also serve as a platform to search for partners for transnational collaboration projects, specially for the EU Sixth Framework Program (NETworks of Excellence and Integrated Projects). "Projects Presentations" and "Call for Partners" presentations proposals are therefore encouraged and welcomed. If you are interested in taking part of this Conference feature, please send us the corresponding form available at the website.
In addition to the "traditional" oral contributed and posters presentation, a Virtual Participation modality has been established for those researchers unable to attend it in person. A limited number of works can be presented in this way. Please refer to the Conference website for details.
If you are interested in taking part of APHYS-2003, please send us your PRE-REGISTRATION FORM (at the main website of the conference) as soon as possible. The pre-registration form is also available through the direct URL http://www.formatex.org/aphys2003/preregistration.htm
Deadline for abstracts submission is April 15th 2003 although we highly recommend you to submit your abstracts as soon as possible to avoid saturation during the days before the deadine (more than 800 researchers are expected to attend this large Applied Physics Conference).
Proceedings Accepted and presented papers will be reviewed for publication in special issues of several international Journals such as Journal of Microscopy, Journal of Non-crystalline Solids, Microelectronics Journal, Physica Scripta, Applied Surface Science, Radiation Protection Dosimetry and Applied Physics A (Materials Science & Processing, to be confirmed). Also a book "Advances in Applied Physics" will be published by an international publisher with those papers accepted for presentation but not suitable for the journal issues. For up-to-date information on publications participating at the Conference as publishers, please visit regularly the Conference website (Proceedings sections).
For any question or suggestion, please do not hesitate to contact us at secretariat-at-formatex.org, or visit www.formatex.org/aphys2003/aphys2003.htm (Bookmark the page!!) We would also appreciate if could disseminate this Call for Papers through your Department or Institution.
We hope to meet you at this exciting and interdisciplinar meeting!
----- Original Message ----- } From: "emlad" {emlad-at-hn.vnn.vn} To: {microscopy-at-sparc5.microscopy.com} Sent: Friday, February 14, 2003 1:14 PM
Dear colleague
This is to inform you that the Call for Abstracts for the forthcoming International Meeting on Applied Physics (APHYS-2003), to be held during October 14-18th 2003 in Badajoz (Spain), is now opened. All the information regarding this interdisciplinary conference can be found at the Conference website
In addition to the regular Scientific Program, several International Workshops will be held as pre-conference events. The following three Workshops are presently confirmed:
1. Workshop on Modern Applied Microscopy in Molecular and Cell Biophysics Research 2. International Interdisciplinar Workshop on Bioengineered Non- crystalline Solids 3. International Workshop on Occupational Radiation Protection
The Conference will be specifically interested in receiving reports on Interdisciplinary researches relating Physics with other Sciences such as Biology, Chemistry, Information Technology, Medicine, etc or relating different Physics areas. In other words, we are specially (but not exclusivelly) interested in reports applying the techniques, the training, and the culture of physics to research areas usually associated with other scientific and engineering disciplines
APHYS-2003 will also serve as a platform to search for partners for transnational collaboration projects, specially for the EU Sixth Framework Program (NETworks of Excellence and Integrated Projects). "Projects Presentations" and "Call for Partners" presentations proposals are therefore encouraged and welcomed. If you are interested in taking part of this Conference feature, please send us the corresponding form available at the website.
In addition to the "traditional" oral contributed and posters presentation, a Virtual Participation modality has been established for those researchers unable to attend it in person. A limited number of works can be presented in this way. Please refer to the Conference website for details.
If you are interested in taking part of APHYS-2003, please send us your PRE-REGISTRATION FORM (at the main website of the conference) as soon as possible. The pre-registration form is also available through the direct URL http://www.formatex.org/aphys2003/preregistration.htm
Deadline for abstracts submission is April 15th 2003 although we highly recommend you to submit your abstracts as soon as possible to avoid saturation during the days before the deadine (more than 800 researchers are expected to attend this large Applied Physics Conference).
Proceedings Accepted and presented papers will be reviewed for publication in special issues of several international Journals such as Journal of Microscopy, Journal of Non-crystalline Solids, Microelectronics Journal, Physica Scripta, Applied Surface Science, Radiation Protection Dosimetry and Applied Physics A (Materials Science & Processing, to be confirmed). Also a book "Advances in Applied Physics" will be published by an international publisher with those papers accepted for presentation but not suitable for the journal issues. For up-to-date information on publications participating at the Conference as publishers, please visit regularly the Conference website (Proceedings sections).
For any question or suggestion, please do not hesitate to contact us at secretariat-at-formatex.org, or visit www.formatex.org/aphys2003/aphys2003.htm (Bookmark the page!!) We would also appreciate if could disseminate this Call for Papers through your Department or Institution.
We hope to meet you at this exciting and interdisciplinar meeting!
Our lab is converting from sending micrographs manually, to sending scanned pictures over e-mail. What type of programs are you using to do this? Are you finding that you need to make adjustments to the pictures before you send them? Also what type of apparatus are you using to show accurate sizing?
Lloyd Willard, Research Associate Department of Diagnostic Medicine/Pathobiology Phone: (785) 532-4420 Electron Microscopy Laboratory Fax: (785) 532-4039 Kansas State University email: lwillard-at-vet.k-state.edu K-208 Mosier Hall web: www.vet.k-state.edu/depts/dmp/personnel/staff/research.htm Manhattan, Ks 66506-5606
} From: WCRGs-at-aol.com } Date: Mon, 17 Feb 2003 15:34:07 EST } Subject: Joel JSM T-300 } To: ListServer-at-sparc5.microscopy.com } Status: } } Hello Listers; } We have a Joel JSM T-300 SEM. This is an old piece of equipment } that we have been working on for a while. We are at the point where } we can get a filament current. There are two problems that we have } run into - 1) when we turn on the accelerating voltage, the filament } lamp lights up immediately. The instruction manual says that the } lamp should not light up until the filament dial is turned to about } the 3:00 o'clock position. This is even with the gun bias at the } full clockwise position. 2) We seem to hear and see (filament } checker spike), some type of spiking occasionally in the area of the } electron gun. This spiking seemed to show up when we started our } gun alignment procedure. } } If anyone has any suggestions on how we might address these problems } they would be much appreciated. } Thanks, } Ronald Obie } Wood Coatings Research Group, Inc. } 336-841-0264
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (ronald.s.najorka-at-intel.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, February 17, 2003 at 15:30:50 ---------------------------------------------------------------------------
Email: ronald.s.najorka-at-intel.com Name: Ron Najorka
Organization: Intel Corp.
Education: Graduate College
Location: Hillsboro,OR,USA
Question: When using the Hitachi s-4700. I was told that if I flashed the tip every time I put a sample in, I would deplete the source to fast. Rather than if I flashed every 8 hours. This is strange because if I flash the tip every 20 minutes for each new sample I get much crisper pictures. Mainly because I am able to work with the aperature and x,y stigs in a much more precise manner.
Hi Tom, I keep a #11 scalpel blade (on the longer small handle) to separate obstreperous slides when I retrieve them from the 70% ethanol into which I immerse them to facilitate the separation. Often, they separate easily after they are wetted.
You could also use 1/2 of a double-edged razor blade in an appropriate holder leaving little of the edge exposed (I recommend a holder lest parts of hands and fingers become mounted along with the section).
Cheers and hope this helps,
Fred Monson Frederick C. Monson, PhD Center for Advanced Scientific Imaging
Mail to: Geology, CASI West Chester University of Pennsylvania Schmucker II Science Center, Room SS024 South Church Street and Rosedale Avenue West Chester, PA, 19383
For help and information only, The CASI houses: An FEI Quanta 400 and Technai 12T, Oxford INCA Energy 400, Tousimis AutoSamdri 815 and Olympus FV-300.
-----Original Message----- } From: Tom Parker [mailto:tparker-at-lacsd.org] Sent: Friday, February 14, 2003 6:55 PM To: microscopy-at-sparc5.microscopy.com
Greetings Microscopers:
Does anyone have a clever easy way to separate new glass slides when you open the box and they are perfectly stuck together. I have heated them in water but that didn't work. I've tapped them on counters, picked at their corners with forceps and fingers....the glass is so clean that they are just "welded" together along their entire surface.
} Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (ronald.s.najorka-at-intel.com) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, } February 17, 2003 at 15:30:50 } --------------------------------------------------------------------------- } } Email: ronald.s.najorka-at-intel.com } Name: Ron Najorka } } Organization: Intel Corp. } } Education: Graduate College } } Location: Hillsboro,OR,USA } } Question: When using the Hitachi s-4700. I was told that if I flashed } the tip every time I put a sample in, I would deplete the source to } fast. Rather than if I flashed every 8 hours. This is strange because } if I flash the tip every 20 minutes for each new sample I get much } crisper pictures. Mainly because I am able to work with the aperature } and x,y stigs in a much more precise manner.
Both parts of this are true. When you flash the tip you are passing a current through it to heat it up (it's normally a cold cathode tip), driving off contaminating molecules. When clean, you get more electrons off with less of an extraction potential, a better signal-to-noise ratio, and crisper pictures. Also true is that whenever you heat up the tip you drive off more material from that tip and cause it to become duller (less pointy), which translates to less resoution over time, plus shorter tip life. The trick is to find the balance, flashing only when it really needs it, usually whatever Hitachi recommends.
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
Depending on which e-mail client you use, the most normal way of sending picture files is via an attachment. These attachments can be encoded using several methods. The most common, I believe, is MIME. Outlook and Eudora offer this method.
One thing to look out for is the total packet size. If your outgoing SMTP server limits total size, you will have to keep your encoded message below that number. MIME messages have somewhere between 25% to 50% overhead. The workaround is to have a public ftp site where files can be dumped but not read.
Whatever the pix looks like before sending, it will look the same at the receiving end--if screen gamma is reasonably the same.
gary g.
At 09:05 AM 2/18/2003, you wrote:
} Our lab is converting from sending micrographs manually, to sending } scanned pictures over e-mail. What type of programs are you using to do } this? Are you finding that you need to make adjustments to the pictures } before you send them? Also what type of apparatus are you using to show } accurate sizing? } } Lloyd Willard, Research Associate } Department of Diagnostic Medicine/Pathobiology Phone: (785) } 532-4420 } Electron Microscopy Laboratory } Fax: (785) 532-4039 } Kansas State University email: lwillard-at-vet.k-state.edu } K-208 Mosier Hall web: } www.vet.k-state.edu/depts/dmp/personnel/staff/research.htm } Manhattan, Ks 66506-5606
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (atthom02-at-louisville.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Tuesday, February 18, 2003 at 22:21:25 ---------------------------------------------------------------------------
Email: atthom02-at-louisville.edu Name: Arun
Organization: Thomas
Education: Graduate College
Location: Louisville, KY
Question: This is Arun Thomas from University of Louisville, Kentucky. We are researching the potential uses of Titanium alloys on medical implant devices. To the Engineering school,a Bausch and Lomb Dynazoom metallograph is donated, but unfortunately it does not have any imaging device with it. So we are interested in getting an attachment to this metallograph. We do not have any contacts for this thing to get done. I would appreciate if you could pass on some contact information regarding the tech support for this machine. Looking forward to hear from you soon, Thanks, Arun
Hi, Some time ago we have solved similar problem by putting the sealed glass slides into common lab ultrasonic cleaner in 70% ethanol. After some time of sonication the glass slides were separated. Maybe it will works for you, too. Best regards Oldrich
+-----------------------------------+ Oldrich Benada Acad. Sci. CR Institute of Microbiology Laboratory of electron microscopy Videnska 1083 CZ - 142 20 Prague 4 - Krc Czech Republic +------------------------------------+ Phone: +420-241062399 Fax: +420-241062347 WEB: http://www.biomed.cas.cz/mbu/lem113/lem.htm
-----Original Message----- } From: Tom Parker [mailto:tparker-at-lacsd.org] Sent: Friday, February 14, 2003 6:55 PM To: microscopy-at-sparc5.microscopy.com
Greetings Microscopers:
Does anyone have a clever easy way to separate new glass slides when you open the box and they are perfectly stuck together. I have heated them in water but that didn't work. I've tapped them on counters, picked at their corners with forceps and fingers....the glass is so clean that they are just "welded" together along their entire surface.
Digital sacanning and transmission of images can be a complex issue, so I am not attempting to be complete here. I will only present some thoughts on digital image scanning and image transmission.
Scanning the images can be done with Adobe Photoshop and several scanners, most of them can be used in combination with a TWAIN driver. Digital images can be sent over the Internet with almost any email program.
Scanning the images at 300 dpi or more is in general good enough for printing purposes afterwards, use about 70 dpi when the images are meant solely for onscreen viewing. Keep in mind that a high resolution of the scanner is not always "hard" but often done by interpolation.
When scanning and transmitting digital images it is important to know what they are meant for. If the images are meant for on-screen viewing, JPEG compression is good enough and a resolution of about 70 dpi (dots per inch) can be used. Care has to be taken if the images are to be used for printing or analysis afterwards. JPEG compression is lossy and introduces artefacts. Tiff with LZW compression is a possible alternative but the file size can be prohibitive.
} } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Our lab is converting from sending micrographs manually, to sending } scanned pictures over e-mail. What type of programs are you using to do } this? Are you finding that you need to make adjustments to the pictures } before you send them? Also what type of apparatus are you using to show } accurate sizing? } } Lloyd Willard, Research Associate } Department of Diagnostic Medicine/Pathobiology Phone: (785) } 532-4420 } Electron Microscopy Laboratory } Fax: (785) 532-4039 } Kansas State University email: lwillard-at-vet.k-state.edu } K-208 Mosier Hall web: } www.vet.k-state.edu/depts/dmp/personnel/staff/research.htm } Manhattan, Ks 66506-5606
Dear Lloyd, We scan pictures (with contrast/brightness adjusted suitably), do any further necessary contrast/brightness adjustments, label them with a micron bar, and then save them as TIFF. They are uploaded to a company server by ftp. We do the image processing using Photoshop but other packages could also be used.
Remember a micron bar is better than a magnification figure because you don't know if the client will blow up the pictures or something, in which case the magnification number becomes worthless, but the micron bar gets blown up with everything else.
As for image formats, TIFF or EPS preserve all information but can be quite big. JPEG is smaller because it is compressed, but the compression is lossy, i.e. not good for archive quality. Photoshop format is nice because layers (text annotations etc.) are preserved but this format may be large and not readable by all programs. In general we use TIFF for this reason, although I preserve a copy in Photoshop if I want to be able to re-edit the annotations later (perhaps for a conference presentation or such like).
Finally, sending by email is problematic as pictures tend to be large files. If you can set up an ftp server at one end or the other, it is much easier.
If you, however, want to just send someone a couple of pics to look at and not for them to use or edit themselves, then putting them into a pdf document is a good solution. Stick them into some programme that combines images and text reasonably well so that you can put a little label underneath each pic. Some such programmes include MS Powerpoint, Deneba Canvas, Adobe Pagemaker. Install Adobe Acrobat (full version) on your computer. Then when you print, you can print to pdfWriter or Distiller and get a pdf file instead of a printout. With suitable setting of the graphics resolution for pdfWriter or Distiller (eg 150 dpi), you get a nice small pdf file with all your pics in. This is great when writing a paper and you want to show the co-authors at other locations your pics without having to send 10 Mb emails. Instead, the pdf file will weigh in at less than 1 Mb with suitable choices for the Distiller or pdfWriter settings.
So,
for archive quality and editable pics --} TIFF,
for quick showing of pics --} all together in one pdf.
Hope this helps
Ian
On Tue, 18 Feb 2003 11:05:47 -0600, EM Lab {Emlab-at-vet.k-state.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Our lab is converting from sending micrographs manually, to sending } scanned pictures over e-mail. What type of programs are you using to do } this? Are you finding that you need to make adjustments to the pictures } before you send them? Also what type of apparatus are you using to show } accurate sizing? } } Lloyd Willard, Research Associate } Department of Diagnostic Medicine/Pathobiology Phone: (785) } 532-4420 } Electron Microscopy Laboratory Fax: } (785) 532-4039 } Kansas State University email: lwillard-at-vet.k-state.edu } K-208 Mosier Hall web: www.vet.k- } state.edu/depts/dmp/personnel/staff/research.htm } Manhattan, Ks 66506-5606 } }
-- Ian MacLaren Technische Universität Darmstadt Material-und Geowissenschaften Petersenstr. 23 64287 Darmstadt Germany
I agree with these comments, having used FE for 20 years. We flash our SEMs last thing at night and then not again until the following evening. If your IP1 vacuum is good enough, this should be OK. Regular flashing does eventually blunt the tip. We flash on 1 and then on 2 and that's it. Watch IP1 when you flash, this gives a good indication of the vacuum status in the gun. It shouldn't go above 2x10-7. You may need a bake since outgassing samples may put H2O or H2 in the gun chamber. This is difficult for the ion pumps to remove.
Barry Lamb
-----Original Message----- } From: Tina Carvalho [mailto:tina-at-pbrc.hawaii.edu] Sent: 18 February 2003 22:30 To: by way of MicroscopyListServer Cc: MicroscopyListserver
} Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (ronald.s.najorka-at-intel.com) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, } February 17, 2003 at 15:30:50 } -------------------------------------------------------------------------- - } } Email: ronald.s.najorka-at-intel.com } Name: Ron Najorka } } Organization: Intel Corp. } } Education: Graduate College } } Location: Hillsboro,OR,USA } } Question: When using the Hitachi s-4700. I was told that if I flashed } the tip every time I put a sample in, I would deplete the source to } fast. Rather than if I flashed every 8 hours. This is strange because } if I flash the tip every 20 minutes for each new sample I get much } crisper pictures. Mainly because I am able to work with the aperature } and x,y stigs in a much more precise manner.
Both parts of this are true. When you flash the tip you are passing a current through it to heat it up (it's normally a cold cathode tip), driving off contaminating molecules. When clean, you get more electrons off with less of an extraction potential, a better signal-to-noise ratio, and crisper pictures. Also true is that whenever you heat up the tip you drive off more material from that tip and cause it to become duller (less pointy), which translates to less resoution over time, plus shorter tip life. The trick is to find the balance, flashing only when it really needs it, usually whatever Hitachi recommends.
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
} Our lab is converting from sending micrographs manually, to sending } scanned pictures over e-mail. What type of programs are you using to do } this?
Are you referring to the image editing software for converting to formats suitable for e-mail? That is, many e-mail servers will not allow for large attachments, and you therefore need to convert to a compressed format like JPEG. JPEG is however a "lossy" format ... make sure your clients are ok with JPEG artifacts. With respect to "what software", most use Photoshop, but many free softwares are available for conversions ... e.g., search the wwweb for "Irfanview" (Windows).
For other image formats (larger file sizes), you may need to set up an FTP server.
} Are you finding that you need to make adjustments to the pictures } before you send them? Also what type of apparatus are you using to show } accurate sizing?
I don't know how you can ^guarantee^ the final print size. But most image formats, including JPEG, can include the print size definition ... such that if you tell your client "If the image is printed as defined, it will be a specific magnification." Personally, I beleieve all images should include a mag reference in the image itself ... ,e.g., a micron bar which will always reference the correct magnification.
Regarding adjustments, each image should be judged independently ... but you should probably assume they will need something ... even if it's only a micron bar, or the image's print size defined. Again, emphasis should be put on Photoshop, or a quantitative and analytical software (e.g., Image Pro Plus, NIH Image or ImageJ). For editing with respect to presentation, Photoshop offers the best user/peer base and choice of excellent texts, as well as being compatible with quantitative plugins.
People have given some good imaging suggestions, so I won't add to that, but I would suggest that you consider not sending the mages directly at all. Many email systems have arbitrary limits on file size and storage per user. Also many folks still seem unable to handle enclosures (!), or get flustered when they try to read there email form a modem connected computer (while at home).
Because of these issues, I typically share SEM images by posting them on my website. Everybody knows how to use their browser, and from my end there are utilities available that will take an entire folder of images and make HTML thumbnail pages that link to the images. If sombeody wants high res images (usually they don't!) I burn a CD and FEDEX it.
--
Michael J. Herron, U of MN, Dept. of Entomology herro001-at-umn.edu 612-624-3212 (lab) St. Paul, MN 55108
We are interested in purchasing an ultra-thin coater for FESEM work. I would appreciate any opinions on types of materials used ie: chrome, platinum, osmium. Also, any preference on manufacturers. We are imaging semiconductors. Our main application for this coater would be cross-sections of semiconductors. Thanks for your input.
Mark Windland Honeywell Plymouth, Minnesota 763-954-2845 mark.j.windland-at-honeywell.com
I wonder if there are standard (or usual) ways for storing setting data from electron microscopes (magnification, working distance, acceleration V, etc.) into the image file itself, such that they can be automatically imported to a database. Some other devices (like digital cameras) automatically use the IPTC or EXIF fields for this.
Any general idea about how preserve and manage these data together with the images will be very welcome.
Martín J. Ramírez División Aracnología Museo Argentino de Ciencias Naturales Av. Angel Gallardo 470 C1405DJR Buenos Aires Argentina tel +54 11 4982-8370 fax +54 11 4982-4494
Question: This is Arun Thomas from University of Louisville, Kentucky. We are researching the potential uses of Titanium alloys on medical implant devices. To the Engineering school,a Bausch and Lomb Dynazoom metallograph is donated, but unfortunately it does not have any imaging device with it. So we are interested in getting an attachment to this metallograph. We do not have any contacts for this thing to get done. I would appreciate if you could pass on some contact information regarding the tech support for this machine. Looking forward to hear from you soon, Thanks, Arun
Arun:
Here's a link that you should investigate:
http://www.diaginc.com/EN/EN.htm
They advertise heavily in Advance Materials & Processes, a magazine for metallurgists. It seems they specialize in digital imaging systems for microscopes.
If you need to get more elaborate in your lab setup, I can pass on some other contacts.
Stu Smalinskas Metallurgist SKF Plymouth, Michigan stu.smalinskas-at-skf.com
__________________________________________________ Do you Yahoo!? Yahoo! Shopping - Send Flowers for Valentine's Day http://shopping.yahoo.com
The first possible explanation is that you have a pretty big leak in the gun which will increase the leakage current from the gun, hence the filament lamp lighting up.
Disassemble the gun column interface and check the "O" rings in that area. Clean the "O" rings by washing in your hands with hot soapy water. When dry gently pull the rings through your fingers, stretching them slightly to check for cracks. These "O" rings have moving surfaces in contact with them so they should be lightly greased.
Good luck
Steve Chapman Senior Consultant Protrain Electron Microscopy Training and Consultancy World Wide Tel +44 (0)1280 816512 Fax +44 (0)1280 814007 www.emcourses.com
----- Original Message ----- } From: "MicroscopyListServer" {zaluzec-at-sparc5.microscopy.com} To: "MicroscopyListserver" {microscopy-at-sparc5.microscopy.com} Sent: Tuesday, February 18, 2003 7:31 AM
Hi,
We have a Philips 201, lately I have been experiencing lighting problems. Different parts of the negative will be more exposed than others--it is not always in the same spot. Today it happened to be a straight line along the bottom of every negative. About 1/2" or less wide---which causes big problems when printing. Fanning corrected most of the problem. My supervisor has watched me saturate the filament and spread the beam to make sure everything was ok---and it was. I always make sure to center the beam and spread evenly before each shot. Interestingly, my supervisor does not seem to experience these lighting problems on her negatives?
It doesn't happen every time, but most of the time, and I have not experienced this problem when using a different scope (philips 401). If the filament is saturated properly and the beam is centered and spread evenly, where are these shadows on the negatives coming from?
HELP---HELP---HELP
Thank You, Lauren Simmerman Pathology Nebraska Health System-EM Lab 402-502-1811
Does anyone have any unused film cartridges for the JOEL Model #JEM-1200EX? We are looking for Kodak or US made. We have cartridges made in Japan but can not get film in US. Any suggestions would be appreciated.
Lorayne E. Ham Scientific Imaging Specialist
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Possibility of ambient light in the room getting to the negatives?
} We have a Philips 201, lately I have been experiencing lighting problems. Different parts of the negative will be more exposed than others--it is not always in the same spot. Today it happened to be a straight line along the bottom of every negative. About 1/2" or less wide---which causes big problems when printing. Fanning corrected most of the problem. } My supervisor has watched me saturate the filament and spread the beam to make sure everything was ok---and it was. I always make sure to center the beam and spread evenly before each shot. Interestingly, my supervisor does not seem to experience these lighting problems on her negatives? } } It doesn't happen every time, but most of the time, and I have not experienced this problem when using a different scope (philips 401). } If the filament is saturated properly and the beam is centered and spread evenly, where are these shadows on the negatives coming from?
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
It seems that you have to open the gun chamber and remove electron gun and then clean a wehnelt , change the filament, if you can see any errosion obviously. To confirm such problem came from dirty gun. 1 After electron gun is removed from the socket, close the gun chamber (without gun) and then evacuate the gun and column until you get ready lamp. Turn on the accelerating voltage, observ that the filament lamp, it should not me light up. 2 you got spike on filament checker, it means that it has a discharge, which is due to dirty wehnelt cap.
I hope that all above will help you to solve the problem Cheers,
Paiboon Nuannin Dept of Physics Faculty of Science Prince of Songkla University Hatyai, Thailand Quoting MicroscopyListServer {zaluzec-at-sparc5.microscopy.com} :
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } From: WCRGs-at-aol.com } } Date: Mon, 17 Feb 2003 15:34:07 EST } } Subject: Joel JSM T-300 } } To: ListServer-at-sparc5.microscopy.com } } Status:ÝÝ } } } } Hello Listers; } } We have a Joel JSM T-300 SEM. This is an old piece of equipment } } that we have been working on for a while. We are at the point where } } we can get a filament current. There are two problems that we have } } run into - 1) when we turn on the accelerating voltage, the filament } } lamp lights up immediately. The instruction manual says that the } } lamp should not light up until the filament dial is turned to about } } the 3:00 o'clock position. This is even with the gun bias at the } } full clockwise position. 2) We seem to hear and see (filament } } checker spike), some type of spiking occasionally in the area of the } } electron gun. This spiking seemed to show up when we started our } } gun alignment procedure. } } } } If anyone has any suggestions on how we might address these problems } } they would be much appreciated. } } Thanks, } } Ronald Obie } } Wood Coatings Research Group, Inc. } } 336-841-0264 } }
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You might investigate whether this is uneven development, rather than uneven illumination. We had some similar problems a year or so ago, that turned out to be due to plugged holes in our nitrogen burst system. This created gradients in the density of the film near the edges where there was insufficient mixing. If you are agitating by hand, it may be something different about the way you and your supervisor do the agitation.
Marie
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Dr. Marie E. Cantino Director, Electron Microscopy Laboratory Associate Professor of Physiology and Neurobiology University of Connecticut Unit 2242 Storrs, CT 06269-2242 Phone: 860-486-3588 Fax: 860-486-6369
Thank you for your timely responses on slides that stick together. I received suggestions that included:
There is no good way to separate them
Try liquid nitrogen dip
Heat them in oven approx 105 C for period of time
Soak them in solvent such as ethanol
Soak them in ethanol and ultrasonic vibrate them
Pry them apart with razor blade or scalpel blade
I tried both the ethanol soak and oven warming on several batches and the oven warming seemed to separate about half of the clumps (most of my clumps are 2-3 slides per).
Suggestions as to the cause included "water weld" from moist packing of slides to oil adhesion from residue of glass cutter.
It was also suggested by one fairly massive user of slides that manufacturers varied tremendously in this problem.
One person suggested the use of "interleaved slides" that have a thin paper insert between each.
Possibly the list could run a poll on the best slides around?
IrfanView is great tiny but powerful viewer. It has nice editing capacity like brightness, contrast, gamma adjustment. It may work with TWAIN scanners also. I used to associate most image formats with that viewer. It handles even big images very well and it's extremely quick! If you need only to scan image, save it in some format and sent it as an attachment in E.mail, you probably may do it with IrfanView (except sending an E.mail). It's also good idea to embed into the image the scale bar. Personally, I prefer to sent to the customers low-res JPEG images with embedded scale bar for viewing purpose only. If customer satisfied with image, I'll send original high-res TIFF upon request. I also prefer to scan images at the highest possible "optical" scanner's resolution (about 1600 dpi, 16 bit) and save this image as a TIFF untouched (for archival purpose), then in Photoshop I reduce resolution and do some adjustments and save a second copy of the file (TIFF) for working purpose (usually 300 dpi). If I do know that client would be interested to see the image, I also create low-res copy of the image in JPEG (72 dpi) at the same time. I usually use macros to do all these tasks automatically. We used to store archival copy of the image on magneto-optical (MO) disk - 5.2 Gb currently per disk, 10 disks on the shelf...
Recently (with help of my daughter) I discover very nice feature in Photoshop-7 (have no idea does it exist in the earlier versions). It's called 'Adjustable Layer' in the "Layers" Menu. It creates layer with predetermined function like brightness/contrast etc. So, you may change parameters actually not changing the original image. You could create a bunch of such 'Adjustable Layers' over single image with different features. You may re-adjust each layer anytime. So, it's very good if you need to adjust your picture for printing etc. Another things I find in Photoshop-7 that Photoshop files actually smaller (30%) than TIFF. It amused me, but this is true at least in a few cases. It seems to me Photoshop-7 handles memory and other stuff differently (much better) than previous versions. Sergey
At 09:35 AM 2/19/03 -0330, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hello Lauren, I had a problem like this a number of years ago with our Philips 300. It turned out to be not the scope, but a problem with agitation during development of the negatives. Do you have the problem if someone else develops your negatives and/or do you have the problem if you develop your supervisor's negatives? Dean Abel Biological Sciences 141 BB University of Iowa Iowa City IA 52242-1324
At 08:05 PM 2/19/2003 +0000, you wrote: } ----------------------------------------------------------------------- } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } ----------------------------------------------------------------------- } Hi, } We have a Philips 201, lately I have been experiencing lighting problems. } Different parts of the negative will be more exposed than others--it is } not always in the same spot. Today it happened to be a straight line along } the bottom of every negative. About 1/2" or less wide---which causes big } problems when printing. Fanning corrected most of the problem. } My supervisor has watched me saturate the filament and spread the beam to } make sure everything was ok---and it was. I always make sure to center the } beam and spread evenly before each shot. Interestingly, my supervisor does } not seem to experience these lighting problems on her negatives? } It doesn't happen every time, but most of the time, and I have not } experienced this problem when using a different scope (philips 401). } If the filament is saturated properly and the beam is centered and spread } evenly, where are these shadows on the negatives coming from? } HELP---HELP---HELP } Thank You, } Lauren Simmerman } Pathology } Nebraska Health System-EM Lab } 402-502-1811
I think we have a very good solution for you with our IBS/e Ion Beam Sputter Deposition and Etching system. The IBS/e, is a thin film deposition system which is designed to improve high resolution electron microscopy imaging by depositing ultra-thin, fine grain metal and carbon films on specimens.
Some characteristics of ion beam sputtered films:
* 5 to 8Å Cr, Ta or W films eliminate charging and increase contrast up to 500kX * Film quantity required is proportional to specimen surface roughness * Films hold down fine particles * Ir Films act as cladding on delicate specimens subject to beam damage * 8Å Cr films can be used when doing EDS without producing X-rays above noise * 80Å Cr support substrates can be produced that are cohesive, amorphous, and smooth
Ion beam sputtered material evolves controllably and repeatably with an energy {25eV. There is no heat or radiation artifacts to decorate specimen detail. Properly deposited films are beyond the resolving power of the highest magnification FESEM image!
The dual axes motion of the stage insures uniform specimen coverage in cracks and crevices of small and large specimens up to 50mm diameter - up to 100mm with the large area stage.
If you want to take the fullest advantage of your FESEM, don't settle for a standard "chromium coater" utilizing planar magnetron technology. It is exciting and revealing to see the benefits of ion beam sputtered films on your samples. We can deposit many different metals and carbon or show you examples of contrast enhancement on various types of specimens from our library of micrographs.
Please contact me for more information or visit our website at www.southbaytech.com. I'll look forward to hearing from you.
DISCLAIMER: South Bay Technology produces equipment and supplies as described above and, therefore, has a vested interest in promoting their use.
Best regards-
David
"Windland, Mark J (MN14)" wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } We are interested in purchasing an ultra-thin coater for FESEM work. I } would appreciate any opinions on types of materials used ie: chrome, } platinum, osmium. Also, any preference on manufacturers. We are imaging } semiconductors. Our main application for this coater would be } cross-sections of semiconductors. } Thanks for your input. } } Mark Windland } Honeywell } Plymouth, Minnesota } 763-954-2845 } mark.j.windland-at-honeywell.com
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Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (random-at-pdx.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, February 19, 2003 at 16:22:20 ---------------------------------------------------------------------------
Email: random-at-pdx.edu Name: Random Diessner
Organization: Portland State University
Education: Undergraduate College
Location: Portland, OR, USA
Question: I work in a lab studying the morphology of various Archaeal viruses and virus-like-particles. There seems to be some controversy in the lab as to whether or not one can carbon coat a grid without a pre-existing support film. My understanding was that one needed a support film such as formvar or butvar for the carbon to be deposited on, other vehemently deny this. HELP!! =)
Mark Windland wrote: ================================================================== We are interested in purchasing an ultra-thin coater for FESEM work. I would appreciate any opinions on types of materials used ie: chrome, platinum, osmium. Also, any preference on manufacturers. We are imaging semiconductors. Our main application for this coater would be cross- sections of semiconductors. Thanks for your input. ================================================================ The Osmium Plasma Coaters, such as the OPC-60 shown on URL http://www.2spi.com/catalog/osmi-coat.html
employs a process that is unique and should never be confused with "sputtering". As a result, this is not a matter of small grain size, it it, from all that one can determine, to be **no** grain size since the nucleation of the growth is on an atomic scale (instead of from "active sites"). The coating itself seems to be amorphous at least down to the level one can make such a determination. A good example of the completely structureless and featureless coating (at extreme magnifications) is on URL http://www.2spi.com/catalog/opc-40.html
In order to demonstrate just how really thin of a layer can be deposited and still have conductivity, see URL http://www.2spi.com/catalog/osmium-plasma-coater-demonstration.html The coating thickness is estimated to be 20 nm, but in any case, one would never get that kind of BSE signal through a high Z layer if it was much more than that.
The total lack of grain size, as well the thinness of the layer, when coupled with the inertness relative to chromium, would make the osmium coatings put down using the OPC units something worth considering. We would be happy to run a demo sample for you anytime, contact me off-line for details for the sample submission.
Disclaimer: SPI Supplies is the distributor for the OPC line of Osmium Plasma Coaters made by Nippon Laser and Electronics in Nagoya, Japan. So quite naturally, it would be in our own interest to see more of these systems being sold!
Chuck
PS: Remember that we are striving to be 100% paperless, therefore there are no paper copies kept of this correspondence. Please be sure to always reply by way of "reply" on your software so that the entire string of correspondence can be kept in one place. ============================================
Charles A. Garber, Ph. D. Ph: 1-610-436-5400 President 1-800-2424-SPI SPI SUPPLIES FAX: 1-610-436-5755 PO BOX 656 e-mail:cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust.Service: spi2spi-at-2spi.com
Look for us! ######################## WWW: http://www.2spi.com ######################## ============================================
Seems a little risky and a waste of liquid N2??????
At 14:34 2003-02-17 -0600, Cheryl Rehfeld - Meyer Instruments, Inc wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Obs/NB New postal/visiting address from July 2002!
Gareth Morgan MPhil MSc FIBMS, Department of Laboratory Medicine (Labmed), Karolinska Institutet, Huddinge Universitetssjukhus, F46 SE 141 86 Stockholm Sweden
OBS! Besöksadress: F-Huset, Forskningsgatan 2 F52, Rum 2.10. Laboratoriet för klinisk patologi och cytologi.
NB! Visiting address: Building F, Research Corridor 2 F52, Room 2.10. Clinical Histo- and Cytopathology Laboratory.
Two other really nice image-viewers are XnView, a free, explorer-like image viewers with great capabilities (like e.g. converting a serie of images from one type to another, e.g. TIFF to JPEG, very fast, a nice overview via tumbnails,...) Download it for free at: http://www.xnview.com
Another VERY interesting program to adjust your photo's is NeatImage. With this program, you increase your sharpness (e.g. take away pixelation). Take a look at http://www.neatimage.com/examples.html. You can download the program, a free demo, but it has enough capabilities at: http://www.neatimage.com/
Really take some time to take a look, it's great!
Sven Terclavers
°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°° Sven Terclavers LM/CLSM Microscopist Center for Transgene Technology and Gene Therapy (CTG) Campus Gasthuisberg K.U.L. O&N Herestraat 49 3000 Leuven Belgium Tel. +32 16 346173 Fax. +32 16 345990 Email: Sven.Terclavers-at-med.kuleuven.ac.be °°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°°
______________ Thursday, February 20, 2003, 1:46:16 AM, you wrote:
SR} ------------------------------------------------------------------------ SR} The Microscopy ListServer -- Sponsor: The Microscopy Society of America SR} To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com SR} On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html SR} -----------------------------------------------------------------------.
SR} IrfanView is great tiny but powerful viewer. It has nice editing capacity SR} like brightness, contrast, gamma adjustment. It may work with TWAIN SR} scanners also. I used to associate most image formats with that SR} viewer. It handles even big images very well and it's extremely quick! If SR} you need only to scan image, save it in some format and sent it as an SR} attachment in E.mail, you probably may do it with IrfanView (except sending SR} an E.mail). It's also good idea to embed into the image the scale bar. SR} Personally, I prefer to sent to the customers low-res JPEG images with SR} embedded scale bar for viewing purpose only. If customer satisfied with SR} image, I'll send original high-res TIFF upon request. I also prefer to scan SR} images at the highest possible "optical" scanner's resolution (about 1600 SR} dpi, 16 bit) and save this image as a TIFF untouched (for archival SR} purpose), then in Photoshop I reduce resolution and do some adjustments and SR} save a second copy of the file (TIFF) for working purpose (usually 300 SR} dpi). If I do know that client would be interested to see the image, I SR} also create low-res copy of the image in JPEG (72 dpi) at the same time. I SR} usually use macros to do all these tasks automatically. We used to store SR} archival copy of the image on magneto-optical (MO) disk - 5.2 Gb currently SR} per disk, 10 disks on the shelf...
SR} Recently (with help of my daughter) I discover very nice feature in SR} Photoshop-7 (have no idea does it exist in the earlier versions). It's SR} called 'Adjustable Layer' in the "Layers" Menu. It creates layer with SR} predetermined function like brightness/contrast etc. So, you may change SR} parameters actually not changing the original image. You could create a SR} bunch of such 'Adjustable Layers' over single image with different SR} features. You may re-adjust each layer anytime. So, it's very good if you SR} need to adjust your picture for printing etc. Another things I find in SR} Photoshop-7 that Photoshop files actually smaller (30%) than TIFF. It SR} amused me, but this is true at least in a few cases. It seems to me SR} Photoshop-7 handles memory and other stuff differently (much better) than SR} previous versions. Sergey
SR} At 09:35 AM 2/19/03 -0330, you wrote: } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
It is possible to evaporate carbon onto a glass slide, float the film onto water and pick the carbon film up on grids much the same as is done with either formvar or butvar. However.... Carbon coated grids prepared in this manner are generally not as strong as formvar or butvar supported films, and thus do not lend themselves to the rigors of negative staining as readily. Preparing such carbon film coated grids is possible. The question is: Why?
Roger Moretz, Ph.D. Dept of Toxicology BI Pharmaceuticals -- Where the world is only slightly less weird than it actually is. } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } 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 (random-at-pdx.edu) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, } February 19, 2003 at 16:22:20 } --------------------------------------------------------------------------- } } Email: random-at-pdx.edu } Name: Random Diessner } } Organization: Portland State University } } Education: Undergraduate College } } Location: Portland, OR, USA } } Question: I work in a lab studying the morphology of various Archaeal } viruses and virus-like-particles. There seems to be some controversy } in the lab as to whether or not one can carbon coat a grid without a } pre-existing support film. My understanding was that one needed a } support film such as formvar or butvar for the carbon to be deposited } on, other vehemently deny this. HELP!! =) } } --------------------------------------------------------------------------- }
Dear All, We have a JEOL 5910LV SEM. Recently, I was testing our peltier cryostage to view wet clay samples. We can freeze up to -25 degrees celsius and set the pressure in the sample chamber to 230Pa at maximum. We did not manage to image any ice or wet material. The samples looked freezedried!
Does anyone has experience with this type of work?
Thanks, Ineke Joosten Netherlands Institute for Cultural Heritage Conservation Research Gabriel Metsustraat 8 1072 EA Amsterdam The Netherlands 00 31 (0)20 3054688/728
We do not have holders for JEOL but do have camera boxes and film cassettes for Philips 400 series microscopes. They are free to a good home as long as you pay shipping. Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building 170 S. University Street West Lafayette, IN 47907
On 2/19/03 3:38 PM, "Lorayne Ham" {Lham-at-snblusa.com} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Does anyone have any unused film cartridges for the JOEL Model #JEM-1200EX? } We are looking for Kodak or US made. We have cartridges made in Japan but } can not get film in US. } Any suggestions would be appreciated. } } Lorayne E. Ham } Scientific Imaging Specialist } } SNBL-USA, Ltd. } 6605 Merrill Creek Parkway } Everett, WA 98203 } (425) 407-0121 ext. 2155 } (425) 407-1122 Fax } email: lham-at-snblusa.com } } Confidentiality Notice: This email, its contents and attachments are } confidential and may contain privileged information. It is intended solely } for the use of addressee(s) only. Any use, copying or disclosure of this } communication or attachments to any other person is expressly prohibited } without written permission of SNBL USA, Ltd. If you receive this message in } error, please notify the sender at SNBL USA, Ltd. immediately by return } e-mail, telephone +1 425 407 0121, or fax +1 425 407 8601. We appreciate } your cooperation. } } } }
Lauren, check your film supply....is it the "new" formulation of Kodak 4489? Your problems sound like the ones reported by people using the new film. 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
Is Os coating durable? I do not use Cr coating because I was getting "disposable" specimens (oxidation was a problem).
Vladimir
} ================================================================ } The Osmium Plasma Coaters, such as the OPC-60 shown on URL } http://www.2spi.com/catalog/osmi-coat.html } } employs a process that is unique and should never be confused with } "sputtering". As a result, this is not a matter of small } grain size, it it, } from all that one can determine, to be **no** grain size since the } nucleation of the growth is on an atomic scale (instead of } from "active } sites"). The coating itself seems to be amorphous at least } down to the } level one can make such a determination. A good example of } the completely } structureless and featureless coating (at extreme } magnifications) is on URL } http://www.2spi.com/catalog/opc-40.html } } In order to demonstrate just how really thin of a layer can } be deposited and } still have conductivity, see URL } http://www.2spi.com/catalog/osmium-plasma-coater-demonstration.html } The coating thickness is estimated to be 20 nm, but in any } case, one would } never get that kind of BSE signal through a high Z layer if } it was much more } than that. } } The total lack of grain size, as well the thinness of the layer, when } coupled with the inertness relative to chromium, would make the osmium } coatings put down using the OPC units something worth } considering. We would } be happy to run a demo sample for you anytime, contact me off-line for } details for the sample submission. } } Disclaimer: SPI Supplies is the distributor for the OPC line } of Osmium } Plasma Coaters made by Nippon Laser and Electronics in } Nagoya, Japan. So } quite naturally, it would be in our own interest to see more of these } systems being sold! } } Chuck } } PS: Remember that we are striving to be 100% paperless, } therefore there } are no paper copies kept of this correspondence. Please be } sure to always } reply by way of "reply" on your software so that the entire string of } correspondence can be kept in one place. } ============================================ } } Charles A. Garber, Ph. D. Ph: 1-610-436-5400 } President 1-800-2424-SPI } SPI SUPPLIES FAX: 1-610-436-5755 } PO BOX 656 e-mail:cgarber-at-2spi.com } West Chester, PA 19381-0656 USA } Cust.Service: spi2spi-at-2spi.com } } Look for us! } ######################## } WWW: http://www.2spi.com } ######################## } ============================================ } } } }
We have an older (model E5000) Polaron sputter coater with a broken switch. So far, I've not had any success in finding a replacement. Does anyone know of a source for Polaron parts?
By the way, the switch is a six-position, two-deck design, with two sweeps per deck. Two-deck, six-position, double-pole switches are easy enough to come by, but I haven't seen any with two sweeps per deck.
Thanks
Kevin L. Macke Research Technician Materials Characterization Facility
phone: (215) 898-4555 fax: (215) 573-0620
Department of Materials Science & Engineering University of Pennsylvania 3231 Walnut Street Philadelphia, PA 19104
There is a Phillips EM300, serial #D997, in excellent working condition that is available for the asking. The scope has been under service contract until last December and will no longer be used by DEC. The instrument has a 35mm, 70 mm and standard photographic plate cameras. There is a mount for a digital camera under the column. Various parts, o-rings, filaments, specimen holders, etc. are also available. The water chiller is not included. This TEM is still using the original mercury pumps. Shipping is up to the individual(s) or institution(s) interested.
Parties interested in this TEM will be considered on a first-come, first-serve basis according to the following priorities:
First priority: Any individual/institution willing to accept the TEM as is, accepting the scope with the mercury pumps and lower vacuum system in place.
Second priority: Any individual/institution willing to accept the TEM as is but with the mercury pumps and lower vacuum system removed.
Third priority: The instrument, as a last resort, will be pieced out to those desiring spare parts for their EM300's.
Interested parties are encouraged to contact me offline.
There are two ways to coat a carbon film on a grid. 1. Coat a freshly cleaved mica, float carbon on water and pick up carbon with a grid from below and blot water off. 2. Coat carbon on previously coated grids (with formvar or other plastic film). If the carbon is thick enough, you can dissolve the film with suitable solvent. Carbon is left on the grids. However, you may use the grids without dissolving the plastic film if it is done with a thin layer of carbon to strengthen it.
AnnFook Yang EM Unit, Eastern Cereal and Oilseed Research Centre, Room 2091, Bldg. 20, Central Experimental Farm, Ottawa, Ontario Canada K1A 0C6
Tel: 1-613-759-1638 Fax: 1-613-759-1701
e-mail: yanga-at-em.agr.ca
} } } by way of MicroscopyListServer {random-at-pdx.edu} 02/20/03 12:54AM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (random-at-pdx.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, February 19, 2003 at 16:22:20 ---------------------------------------------------------------------------
Email: random-at-pdx.edu Name: Random Diessner
Organization: Portland State University
Education: Undergraduate College
Location: Portland, OR, USA
Question: I work in a lab studying the morphology of various Archaeal viruses and virus-like-particles. There seems to be some controversy in the lab as to whether or not one can carbon coat a grid without a pre-existing support film. My understanding was that one needed a support film such as formvar or butvar for the carbon to be deposited on, other vehemently deny this. HELP!! =)
I have tried two variations on the theme. I have used a paper punch to punch out rounds of freshly cleaved mica, stuck one edge of each round onto a clean glass slide with double stick tape, and evaporated carbon onto the slide. Score around the edges of the coated mica, or make a tic-tac-toe grid on each with a needle, leaving the center square large enogh for a grid, then float the films off the mica (one by one) onto water. Place a grid on the film and pick up. My favorite way to pick them up (and which I also use for making Formvar-coated grids) is to come down on top of them with a piece of Parafilm, then lift the Parafilm off. The films seem to float off the mica pieces easier than off a slide, at least in my hands. I've made some pretty sturdy and thin films this way - mostly to image nanoparticles.
Alternatively, I have evaporated carbon onto Formvar-coated grids, stuck them onto a slide as above, then dissolved away the plastic film. With uneven success, I must admit. Right now I can't remember what solvent(s) worked the best, and I often ended up with shreds of Formvar remaining on the grid. However, in these cases I still had enough pure carbon areas that I could easily image proteins and particles.
The pure carbon films do allow much better resolution and contrast than the Formvar or Butvar, but are certainly more hassle!
Aloha, Tina
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
Brown University announces an opening for a microscopist/manager in the Electron Microscopy Facility in the Center for Advanced Materials Research (CAMR) at Brown University. This central facility serves users in Engineering, Physics, Geology, and Chemistry, as well as visitors from academia and industry. This user-operated facility contains modern electron imaging tools including TEM (JEOL 2010, EM420) and SEM (LEO 1530VP, JEOL 845) and also includes state-of-the-art optical and scanning probe microscopy equipment. The ideal candidate will have experience in the use of transmission electron microscopy for materials research and will teach a graduate level lab course in this area. The facilities manager oversees the daily operation of the facility, trains new users, and works with faculty on sponsored research projects. Other responsibilities of this position include representing the facility in dealings with equipment and service vendors, and troubleshooting sophisticated microscopy and sample preparation equipment. The education and experience of the successful candidate should be equivalent to a Masters level degree in materials science (or a closely related field) and include five to seven years of practical experience. Exceptional candidates with clearly demonstrated expertise in the required areas will also be considered.
Contact:
Professor David C. Paine Brown University Division of Engineering, Box D 182 Hope Street Providence, RI02912
On Wednesday, February 19, 2003, at 09:54 PM, by way of MicroscopyListServer wrote:
} Question: I work in a lab studying the morphology of various Archaeal } viruses and virus-like-particles. There seems to be some controversy } in the lab as to whether or not one can carbon coat a grid without a } pre-existing support film. My understanding was that one needed a } support film such as formvar or butvar for the carbon to be deposited } on, other vehemently deny this. HELP!! =) } Dear Random, Since we cryo-EM folks routinely make holey carbon grids by evaporating carbon onto a plastic film with ~1 - ~10 micrometer holes in it, and, since there is no carbon where there were holes, I would definitely say that you will need a continuous support film on your grid in order to get carbon across the grid openings. If the grid is a high enough mesh, you could dissolve away the formvar, and the carbon would stay intact, but this will not be possible for larger mesh grids. Yours, Bill Tivol 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
You have to have atmosphere of water vapor in a specimen chamber, not air (I do not know if a JEOL 5910LV SEM has this capability).
Even if water vapor is injected in the chamber, it is often not enough during initial pumping, and atmosphere could became too dry. It is recommended to put some additional water droplets into the chamber, preferably close to specimen.
Vladimir
Vladimir M. Dusevich, Ph.D. Electron Microscope Lab Manager 3127 School of Dentistry 650 E. 25th Street Kansas City, MO 64108-2784
} ---------. } } } Dear All, } We have a JEOL 5910LV SEM. Recently, I was testing our } peltier cryostage to } view wet clay samples. We can freeze up to -25 degrees } celsius and set the } pressure in the sample chamber to 230Pa at maximum. We did } not manage to } image any ice or wet material. The samples looked freezedried! } } Does anyone has experience with this type of work? } } Thanks, } Ineke Joosten } Netherlands Institute for Cultural Heritage } Conservation Research } Gabriel Metsustraat 8 } 1072 EA Amsterdam } The Netherlands } 00 31 (0)20 3054688/728 } } } }
While we're on the subject, does anyone know of a freeware DVD player program?
cheers
rtch
} } } Two other really nice image-viewers are XnView, a free, explorer-like } image viewers with great capabilities (like e.g. converting a serie of } images from one type to another, e.g. TIFF to JPEG, very fast, a nice } overview via tumbnails,...) Download it for free at: } http://www.xnview.com } } Another VERY interesting program to adjust your photo's is NeatImage. } With this program, you increase your sharpness (e.g. take away } pixelation). Take a look at http://www.neatimage.com/examples.html. } You can download the program, a free demo, but it has enough } capabilities at: http://www.neatimage.com/ } } Really take some time to take a look, it's great! } } Sven Terclavers } } } } } } SR} IrfanView is great tiny but powerful viewer. It has nice editing } capacity SR} like brightness, contrast, gamma adjustment. It may work } with TWAIN SR} scanners also. I used to associate most image formats } with that SR} viewer. It handles even big images very well and it's } extremely quick! If SR} you need only to scan image, save it in some } format and sent it as an SR} attachment in E.mail, you probably may do } it with IrfanView (except sending SR} an E.mail). It's also good idea } to embed into the image the scale bar. SR} Personally, I prefer to } sent to the customers low-res JPEG images with SR} embedded scale bar } for viewing purpose only. If customer satisfied with SR} image, I'll } send original high-res TIFF upon request. I also prefer to scan SR} } images at the highest possible "optical" scanner's resolution (about } 1600 SR} dpi, 16 bit) and save this image as a TIFF untouched (for } archival SR} purpose), then in Photoshop I reduce resolution and do } some adjustments and SR} save a second copy of the file (TIFF) for } working purpose (usually 300 SR} dpi). If I do know that client would } be interested to see the image, I SR} also create low-res copy of the } image in JPEG (72 dpi) at the same time. I SR} usually use macros to } do all these tasks automatically. We used to store SR} archival copy } of the image on magneto-optical (MO) disk - 5.2 Gb currently SR} per } disk, 10 disks on the shelf... }
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
Dear Random, I prepare carbon films by evaporating carbon onto a collodion-covered grid. Then the collodion is dissolved by putting the grid on a chloroform-soaked filter paper stack for 48 hours ("Jaffe washer"). This leaves an amorphous, 20 to 30 nanometer-thick carbon film adhered to the grid. I find these films more robust than the formvar films in my 200kV TEM because they are conductive. I use them to study small particles. Mary ----- Original Message ----- } From: "by way of MicroscopyListServer" {random-at-pdx.edu} To: "MicroscopyListserver" {microscopy-at-sparc5.microscopy.com} Sent: Wednesday, February 19, 2003 9:54 PM
Does any electron microscopist out there in cyber space have a single tilt holder (PW6596) for 3.00 mm grids from a machine they are decommissioning. We would be prepared to pay for shipping and an agreed price.
Hoping someone can help us
Terry Robertson
Dr Terry A Robertson (PhD) Senior Research Fellow School of Surgery and Pathology Division of Pathology University of Western Australia Nedlands Australia 6907 Phone (61) 8 93462935 Mobile 0403025440 Fax (61) 8 93462891 email terryr-at-cyllene.uwa.edu.au
Ineke Joosten wrote: =============================================================== We have a JEOL 5910LV SEM. Recently, I was testing our peltier cryostage to view wet clay samples. We can freeze up to -25 degrees celsius and set the pressure in the sample chamber to 230Pa at maximum. We did not manage to image any ice or wet material. The samples looked freezedried!
Does anyone has experience with this type of work? ================================================================ Once you get above the range of 55-60°C, the sublimation rate of ice becomes considerable. Below that temperature range the rate is very slow. Since you are in the fast sublimation rate range, it would seem that the ice disappeared on you and that is why you are getting the appearance you are seeing.
You would have to be lower in temperature to keep the ice from subliming quickly.
Charles A. Garber, Ph. D. Ph: 1-610-436-5400 President 1-800-2424-SPI SPI SUPPLIES FAX: 1-610-436-5755 PO BOX 656 e-mail:cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust.Service: spi2spi-at-2spi.com
Look for us! ######################## WWW: http://www.2spi.com ######################## ============================================
My statement should have read: ============================ Once you get above the range of -55 to -60°C, the sublimation rate of ice becomes considerable. Below that temperature range the rate is very slow. Since you are in the fast sublimation rate range, it would seem that the ice disappeared on you and that is why you are getting the appearance you are seeing. ============================= In my original posting I said "55-60°C" . Sorry for not better proof- reading.
My recollection of the discription of elemental osmium in the CRC handbook discribes it as having a discernable oder due to the oxidation of metallic Os to the tetroxide... And OsO4 has enough of a vapor pressure to be used as a heavy-metal stain/fixitive in biological TEM.
Ben Simkin (simkin-at-egr.msu.edu) Michigan State University, dpt. Chemical Engineering and Materials Science
On Thu, 20 Feb 2003, Dusevich, Vladimir wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Is Os coating durable? } I do not use Cr coating because I was getting } "disposable" specimens (oxidation was a problem). } } Vladimir } } } ================================================================ } } The Osmium Plasma Coaters, such as the OPC-60 shown on URL } } http://www.2spi.com/catalog/osmi-coat.html } } } } employs a process that is unique and should never be confused with } } "sputtering". As a result, this is not a matter of small } } grain size, it it, } } from all that one can determine, to be **no** grain size since the } } nucleation of the growth is on an atomic scale (instead of } } from "active } } sites"). The coating itself seems to be amorphous at least } } down to the } } level one can make such a determination. A good example of } } the completely } } structureless and featureless coating (at extreme } } magnifications) is on URL } } http://www.2spi.com/catalog/opc-40.html } } } } In order to demonstrate just how really thin of a layer can } } be deposited and } } still have conductivity, see URL } } http://www.2spi.com/catalog/osmium-plasma-coater-demonstration.html } } The coating thickness is estimated to be 20 nm, but in any } } case, one would } } never get that kind of BSE signal through a high Z layer if } } it was much more } } than that. } } } } The total lack of grain size, as well the thinness of the layer, when } } coupled with the inertness relative to chromium, would make the osmium } } coatings put down using the OPC units something worth } } considering. We would } } be happy to run a demo sample for you anytime, contact me off-line for } } details for the sample submission. } } } } Disclaimer: SPI Supplies is the distributor for the OPC line } } of Osmium } } Plasma Coaters made by Nippon Laser and Electronics in } } Nagoya, Japan. So } } quite naturally, it would be in our own interest to see more of these } } systems being sold! } } } } Chuck } } } } PS: Remember that we are striving to be 100% paperless, } } therefore there } } are no paper copies kept of this correspondence. Please be } } sure to always } } reply by way of "reply" on your software so that the entire string of } } correspondence can be kept in one place. } } ============================================ } } } } Charles A. Garber, Ph. D. Ph: 1-610-436-5400 } } President 1-800-2424-SPI } } SPI SUPPLIES FAX: 1-610-436-5755 } } PO BOX 656 e-mail:cgarber-at-2spi.com } } West Chester, PA 19381-0656 USA } } Cust.Service: spi2spi-at-2spi.com } } } } Look for us! } } ######################## } } WWW: http://www.2spi.com } } ######################## } } ============================================ } } } } } } } } }
This is in response to your question about carbon films on grids. You received a response involving carbon evaporated onto cleaved mica. Years ago, we used to buy such 10 NM carbon foils on mica from the Arizona Carbon Foil Co., now called ACF Metals. They were beautifully made but not cheap, as I recall. They will essentially make any thickness carbon foil you could ever want. } From the web, I see they are still in business.
http://www.techexpo.com/WWW/acf-metals/page1.html
http://www.techexpo.com/firms/acf-metl.html
This bottom link still says they supply EM substrates.
Disclaimer: I don't work for ACF Co. or ACF-Metals.
I hope this helps.
Paul Beauregard Senior Research Associate PPG Industries Monroeville Technical Center 440 College Park Drive Monroeville, PA 15146 724-325-5131 pabeauregard-at-ppg.com
Chuck - you could subtract 30oC to your figures. It rather depends what you mean by "slow". My rule of thumb over many years of LTSEM with Cambridge 250/Emscope SP2000 and Hitachi 4700/Gatan Alto has been to use -80oC as the standard etching temperature. From the point of view of LTSEM specimens etching is uncontrollably fast at -60oC, conveniently rapid at -80, slower and more controllable at -90, but is observable down to -100oC, probably lower, since water ice has low but measurable vapour pressure beyond -100oC. If ice must be observed at these temperatures the chamber atmosphere must contain water vapour at a partial pressure equilibrated to the vapour pressure of water above the ice. This can probably only be achieved in ESEM.
Ineke - Whatever the temperature of your cryostage, it is a major technical problem, and one we are grappling with currently, to know how to get an ice specimen into a cryoSEM without either removing ice from its surface or adding ice to its surface. Anyone got an answer to this?
Best wishes Chris
} -- [ From: Garber, Charles A. * EMC.Ver #3.1 ] -- } } Ineke Joosten wrote: } =============================================================== } We have a JEOL 5910LV SEM. Recently, I was testing our peltier cryostage to } view wet clay samples. We can freeze up to -25 degrees celsius and set the } pressure in the sample chamber to 230Pa at maximum. We did not manage to } image any ice or wet material. The samples looked freezedried! } } Does anyone has experience with this type of work? } ================================================================ } Once you get above the range of 55-60°C, the sublimation rate of ice becomes } considerable. Below that temperature range the rate is very slow. Since } you are in the fast sublimation rate range, it would seem that the ice } disappeared on you and that is why you are getting the appearance you are } seeing. } } You would have to be lower in temperature to keep the ice from subliming } quickly. } } Chuck } ============================================ } } Charles A. Garber, Ph. D. Ph: 1-610-436-5400 } President 1-800-2424-SPI } SPI SUPPLIES FAX: 1-610-436-5755 } PO BOX 656 e-mail:cgarber-at-2spi.com } West Chester, PA 19381-0656 USA } Cust.Service: spi2spi-at-2spi.com } } Look for us! } ######################## } WWW: http://www.2spi.com } ######################## } ============================================ } } }
Vladimir Dusevich wrote: ====================================================== Is Os coating durable? I do not use Cr coating because I was getting "disposable" specimens (oxidation was a problem). ======================================================= The osmium metal coating is "durable", indeed relative to chromium, as you suggest, it is inert. After all, it is a precious group metal. Researchers in Japan, where a number of these systems have been installed and used for some years, seem to find that the shelf life for a coated sample is like it would be for gold. Now we have not been able to verify that yet ourselves but specimens coated two years ago by us seem unchanged (when viewed in a conventional non-FESEM instrument, from the way they looked when originally coated.
In order for the Os metal coating to become unstable, it would have to be subjected to some kind of oxidizing agent (and it could be converted back first to the dioxide and then to the tetroxide and that obviously would not be a good thing) but samples sleeping in storage boxes tend to not get exposed to oxidizing agents.......but admittedly, if one was coating particles of sodium periodate, for example, something we have not done, then perhaps one could speculate about its long term stability. But again this is not something we have done.
Chuck
============================================
Charles A. Garber, Ph. D. Ph: 1-610-436-5400 President 1-800-2424-SPI SPI SUPPLIES FAX: 1-610-436-5755 PO BOX 656 e-mail:cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust.Service: spi2spi-at-2spi.com
Look for us! ######################## WWW: http://www.2spi.com ######################## ============================================
Energy Beam Sciences is the exclusive representative for the Polaron Range in the US. We can provide a full range of new instruments as well as parts and service for older ones. you can reach us at (800)992-9037, by email at ebs-at-ebsciences.com or on the web at www.ebsciences.com.
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Michael R. Nesta General Manager Energy Beam Sciences, Inc. 11 Bowles Road Agawam, MA 01001-2925 Tel: (413) 786-9322 Fax: (413) 789-2786 "Adding Brilliance to Your Vision"
-----Original Message----- } From: Kevin Macke [mailto:macke-at-lrsm.upenn.edu] Sent: Thursday, February 20, 2003 11:56 AM To: Microscopy-at-sparc5.microscopy.com
We have an older (model E5000) Polaron sputter coater with a broken switch. So far, I've not had any success in finding a replacement. Does anyone know of a source for Polaron parts?
By the way, the switch is a six-position, two-deck design, with two sweeps per deck. Two-deck, six-position, double-pole switches are easy enough to come by, but I haven't seen any with two sweeps per deck.
Thanks
Kevin L. Macke Research Technician Materials Characterization Facility
phone: (215) 898-4555 fax: (215) 573-0620
Department of Materials Science & Engineering University of Pennsylvania 3231 Walnut Street Philadelphia, PA 19104
Random - You should be able to coat freshly cleaved mica with carbon, float the carbon off onto water, let the water out so the carbon floats down onto grids that you have placed on a wire mesh. You should be able to use up to 400 mesh grids. The smaller the mesh the less breakage.
Please feel free to contact me off line or by phone if you have any questions.
ML -- Mei Lie Wong Electron Microscope Laboratory Department of Biochemistry HHMI-UCSF Ph. 415-476-4441 Fax 415-476-1902 http://util.ucsf.edu/agard/wong/index.html email wong-at-msg.ucsf.edu
We routinely C coat glass slides, float off on water and pick up on 200 mesh grids. The C is coated to a light grey color and works fine for, e.g., 100 Å thick polyethylene single crystals; they may, however, be too thick for some biological samples These will often be dried down on the C coated slides and shadowed before floating. The C is scratched with the point of a tweezers to ca 1/8 in squares before floating. These are then picked up with a grid held in the tweezers, coming up at an angle so the carbon catches on one edge first. If the C layer sticks to the grid, breathing on it after scratching or storing it under an evaporator dish with a small amount of water helps. At times we will also use cover slips as the initial substrate, floating them on ca. 1% HF. There is no way, however, that a C film can be directly deposited on the empty holes of a grid.
--
Phillip H. Geil; Ph. 217-333-0149 Fax 217-333-2736 Department of Materials Science and Engineering University of Illinois 1304 W. Green St. Urbana, IL 61801
Hi all, I'm currently using and Epson 1280 to print photos and it has been adequate for my needs as far as image quality goes but way too slow for printing large numbers of photo quality prints. These past couple of weeks I'm had the need to print large numbers of 8x10 prints FAST! Management wants to purchase a printer that can output a photo quality color print in a minute or less (preferably less!). I just tested a Xerox/Tectronix 6200. The quality was less than the Epson but much faster.
Any recommendations for a fast photo quality printer for digital images? We generally have image files in the 10's of MB and can download to the network 500MB or more worth of photos for a single run.
TIA Damian Neuberger Senior Research Scientist Baxter Healthcare Corp. damian_neuberger-at-baxter.com Tel: 847.270.5888 Fax: 847.270.5897
1) when people want a picture that looks like a photographic print, I use a dye-sub (Kodak). Very slow, software isn;t all that great, and the consumables are very expensive (as was the printer itself, once upon a time). This is used for less than 1% of the work done, and when the printer finally dies it won't be replaced.
2) an Epson outfitted with the Piezographics inks and software, used for about 5% of the printing, produces better-than-photo-quality grey scale prints but is very slow, and also moderately costly per print because of the need for special coated papers. Using grey-scale inks produces fabulous results, but of course this doesn't help much with color (ink jet color prints are so-so; using archival pigmented inks instead of dyes causes some funny color shifts, but the dyes degrade badly with light, heat or humidity).
3) The other 95% is done on a Minolta QMS 3100C laser printer (1200dpi). It is extremely fast, networked to half a dozen computers, uses standard paper, and produces results roughly equivalent to a good magazine or book print. Laser printers have gotten very good for color work (not so wonderful for grey scale). It sounds as though this would fit your needs well. Software is excellent - has built in ICC curves and produces accurate color renditions.
John Russ
======
In a message dated 2/21/03 6:04:30 PM, neuberger1234-at-attbi.com writes:
} I'm currently using and Epson 1280 to print photos and it has been adequate } for my needs as far as image quality goes but way too slow for printing } large numbers of photo quality prints. These past couple of weeks I'm } had } the need to print large numbers of 8x10 prints FAST! Management wants } to } purchase a printer that can output a photo quality color print in a minute } or less (preferably less!). I just tested a Xerox/Tectronix 6200. The } quality was less than the Epson but much faster. } } Any recommendations for a fast photo quality printer for digital images? } We } generally have image files in the 10's of MB and can download to the network } 500MB or more worth of photos for a single run.
There is free open source software: http://gallery.menalto.com/
It allows you to set up image albums very fast and efficient, with differientiated user permissions.
That way, you can avoid e-mailing images altogether, and your customers, clients *or* students can get images at the resolutions they want. You can also ban certain users from downloading full size images for whatever reasons.
Wolf Schweitzer
A "little" more detailed description; http://www.swisswuff.ch/pnphoenix721/html/ modules.php?op=modload&name=News&file=article&sid=26&mode=thread&order=0 &thold=0
On Mittwoch, Februar 19, 2003, at 02:05 Uhr, michael shaffer wrote:
} ----------------------------------------------------------------------- } - } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ----------------------------------------------------------------------- } . } } } Lloyd Willard writes ... } } } Our lab is converting from sending micrographs manually, to sending } } scanned pictures over e-mail. What type of programs are you using to } } do } } this? } } Are you referring to the image editing software for converting to } formats } suitable for e-mail? That is, many e-mail servers will not allow for } large } attachments, and you therefore need to convert to a compressed format } like } JPEG. JPEG is however a "lossy" format ... make sure your clients are } ok } with JPEG artifacts. With respect to "what software", most use } Photoshop, } but many free softwares are available for conversions ... e.g., search } the } wwweb for "Irfanview" (Windows). } } For other image formats (larger file sizes), you may need to set up } an FTP } server. } } } Are you finding that you need to make adjustments to the pictures } } before you send them? Also what type of apparatus are you using to } } show } } accurate sizing? } } I don't know how you can ^guarantee^ the final print size. But most } image } formats, including JPEG, can include the print size definition ... } such that } if you tell your client "If the image is printed as defined, it will } be a } specific magnification." Personally, I beleieve all images should } include a } mag reference in the image itself ... ,e.g., a micron bar which will } always } reference the correct magnification. } } Regarding adjustments, each image should be judged independently ... } but } you should probably assume they will need something ... even if it's } only a } micron bar, or the image's print size defined. Again, emphasis should } be } put on Photoshop, or a quantitative and analytical software (e.g., } Image Pro } Plus, NIH Image or ImageJ). For editing with respect to presentation, } Photoshop offers the best user/peer base and choice of excellent } texts, as } well as being compatible with quantitative plugins. } } hth & cheerios ... shAf :o) } Avalon Peninsula, Newfoundland } www.micro-investigations.com (in progress) } }
} } } Hello All: } } } We are contemplating upgrading or trading-in our SEMs } } } including Hitachi S900. Is anyone out there with a } } } functional Hitachi S900 with EDX on it or at least memories of such a } } } system functioning anywhere, who would be willing to discuss its } } } pro-s and con-s ? } } } Marek. } } } } name MAREK MALECKI } building 1052 ANSCI BUILDING } department ANIMAL SCIENCE } division COLLEGE OF AGRICULTURAL & LIFE SCIENCES } email mmalecki-at-wisc.edu } phone (608) 262-0816 } title PROFESSOR } work-email mmalecki-at-wisc.edu } work_address 1675 OBSERVATORY DR MADISON WI 53706 } } } }
I recently looked through a number of recipes for mounting media that were posted a while back. I was looking for something that hardened so that I did not need to use nail polish to seal coverslip edges. Many recipes use Mowiol but I am confused by what role Mowiol plays in mounting media. I thought its job was to harden upon exposure to air so that the coverslip was sealed. However, the Calbiochem catalog lists it as an antifade reagent. If it is not a hardening agent, then what does that job? Also, the protocols for mounting media using polyvinyl alcohol look identical to that for Mowiol. Is Mowiol a trade name of pva? Thanks- Dave --
Dr. David Knecht Department of Molecular and Cell Biology University of Connecticut 91 N. Eagleville Rd. U-3125 Storrs, CT 06269-3125 knecht-at-uconn.edu 860-486-2200 860-486-4331 (fax) home page: http://www.sp.uconn.edu/~mcbstaff/knecht/knecht.html
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I was just going to ask at what stage of freezing on the Peltier stage do you actually press the evacuate button and how thick were your specimens. If I am working with anything that I think is likely to lose water easily I wait until the sample has passed about -5 deg C before evacuating to between 70 or ~200 Pa. A thick sample of course may not reach the stage temperature as quickly.
But the use of extra water in the chamber is interesting. Does this not play havoc with the vacuum pumps even on a high pressure system - eg a lot extra gas ballasting or special water traps?
Malcolm
Malcolm Haswell e.m. unit School of Health, Natural & Social Sciences University of Sunderland UK
----- Original Message ----- } From: "Dusevich, Vladimir" {dusevichv-at-umkc.edu}
Hi all, I am considering poster printers and wanted input from the multiple users out there that print large photos on poster printers. I am considering both HP and EPSON. Do you have any comments that I should know about. Thanks in advance, Robert
Dr. Robert K. Pope Indiana University Department of Biology 1700 Mishawaka Avenue South Bend, IN 46634 ropope-at-iusb.edu
I have run into a problem when I set a computer monitor next to my Amray 1830I monitor. The computer monitor shows a moving scan line of the SEM monitor and the scan line is gone when the TV is off. My concern is that the computer monitor would get damaged with time. Did anybody have that kind of problem? Is there a B&W TV monitor that would not cause this kind of effect? Any other suggestions. I would prefer to keep the computer monitor next to the TV.
I would like to hear from any vendors out there who have digital camera systems for mounting on TEM. We're looking at replacing our current system mounted on our Philips 410LS TEM. We will probably be most interested in a side mounted system. My bosses requirements are largest possible viewing field, at highest possible resolution, and a real time viewing of image on the monitor during scope operation. I lack experience in this area so any and all advice, information etc. would be greatly appreciated from anyone. Thanks.
Tom Bargar Electron Microscopy Core Research Facility Dept. of Cell Biology and Anatomy 986395 Nebraska Medical Center Omaha, NE 68198-6395
I have heard John McKenzie make the following suggestion at least two times at his seminars. I haven't tried it yet, but I can't argue with the logic.
He suggested that instead of purchasing a single fast, high-end printer, purchase multiple cheap printers and hook them up in parallel. If the cheap printers are 10x slower than the fast one, just hook up 10x as many on a print server. I haven't tracked the Codonics prices lately, but they were running $10K a few years ago. That would buy twenty Epson 1820s. However, I would hold back some and spend some money on the print server that would distribute jobs to the printers. I believe Windows NT (2000, XP-Pro) has such an ability to distribute jobs among similar printers. I just have never had the occasion to try it.
Regarding image sizes, I would seriously investigate this matter. I rather doubt that an Epson could render a 2000-pixel wide image so that it appeared much better than a 1200-pixel wide image when printed at 8x10 inches. It depends on the capabilities of the printer, but a 1200-dpi printer doesn't render 1200 pixels per inch - it probably can only render a tenth of that or about 120 pixels per inch. If so, all the extra pixels in the image are for naught - at least when it comes to printing.
I suspect (but cannot yet prove) that some print drivers are probably better than others are working with this limitation. There is no sense in sending data to the printer in excess of what can be rendered. However, I think some printer drivers are written to send all the data over anyway and let the printer sort it out. But that takes more time to communicate all that data and takes time at the printer to realize it is superfluous. I think a better driver would know the printer capabilities and only send such a data stream as would be useful.
If someone has experience with these matters, I would appreciate hearing the outcome of their experiments.
Warren
At 04:28 PM 2/21/03 -0600, you wrote: } Hi all, } I'm currently using and Epson 1280 to print photos and it has been adequate } for my needs as far as image quality goes but way too slow for printing } large numbers of photo quality prints. These past couple of weeks I'm had } the need to print large numbers of 8x10 prints FAST! Management wants to } purchase a printer that can output a photo quality color print in a minute } or less (preferably less!). I just tested a Xerox/Tectronix 6200. The } quality was less than the Epson but much faster. } } Any recommendations for a fast photo quality printer for digital images? We } generally have image files in the 10's of MB and can download to the network } 500MB or more worth of photos for a single run. } } TIA } Damian Neuberger } Senior Research Scientist } Baxter Healthcare Corp. } damian_neuberger-at-baxter.com } Tel: 847.270.5888 } Fax: 847.270.5897
------------------------------------------- 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
We have a client looking at individual muscle fibers which need to be fixed and embedded for immunolabeling and which will require longitudinal sections along at least of portion of their miniscule lengths. These things are so tiny that they are barely visible with the naked eye in the fixative. They almost look like little dust motes, if we can see them at all.
Immuno means no osmium, so the problem arises about how we find them once they're in the resin (probably LR White), let alone position them precisely enough to do long sections.
Ideas we're considering include: 1) tying the fibers along a tiny section of dark thread so we can at least see the thread; 2) staining them with an LM stain, such as toluidine blue, before embedding them; and 3) settling them down onto poly-l-lysine coated Thermonox cover slips to stabilize them, then scribing the coverslips and/or using an LM stain.
We are trying to avoid pre-embedding labeling at the client's request, but that will be an option if all else fails. Then we can osmicate following the labeling process.
Questions: Has anyone ever tried adhering muscle fibers to poly lysine coverslips? If so, is the attachment stable enough to get through the processing/embedding process? Is the use of an LM stain practical, or does this introduce a lot of contamination at the EM level? Does toluidine blue penetrate past the fiber membrane? (I've never looked at LM stained materials in a TEM before.)
We're trying to get an idea of what might work before we start, since there is a substantial amount of time-consuming dissection, etc., involved.
Thanks much for any ideas and thoughts.
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We're the Fun Core! 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/
"The statements and opinions expressed here by Gary M. Brown represent neither those of ExxonMobil Corporation nor its affiliates."
Damian,
John Mackenzie (North Carolina State) suggested simultaneous printing of images to several Epson printers in parallel. The printers provide great quality images and are dirt-cheap. Several of these in the lab allows overnight printing of a large number of images. I support this approach since it provides (1) the quality, (2) the price, and (3) quantity of images that you need. Remember that what your management wants and what they (you) need are often very different. It is your job to convince them that your approach provides what they want.
Good luck,
Gary M. Brown ExxonMobil Chemical Company Baytown Polymers Center 5200 Bayway Drive Baytown, Texas 77520-2101 phone: (281) 834-2387 fax: (281) 834-2395 e-mail: Gary.M.Brown-at-ExxonMobil.com
"Damian Neuberger" {neuberger1234-at-att To: MicroscopyListserver bi.com} (by way of {microscopy-at-sparc5.microscopy.com} MicroscopyListServ cc: er) Subject: Printer recommendation
02/21/03 04:28 PM
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi all, I'm currently using and Epson 1280 to print photos and it has been adequate for my needs as far as image quality goes but way too slow for printing large numbers of photo quality prints. These past couple of weeks I'm had the need to print large numbers of 8x10 prints FAST! Management wants to purchase a printer that can output a photo quality color print in a minute or less (preferably less!). I just tested a Xerox/Tectronix 6200. The quality was less than the Epson but much faster.
Any recommendations for a fast photo quality printer for digital images? We generally have image files in the 10's of MB and can download to the network 500MB or more worth of photos for a single run.
TIA Damian Neuberger Senior Research Scientist Baxter Healthcare Corp. damian_neuberger-at-baxter.com Tel: 847.270.5888 Fax: 847.270.5897
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (pgan-at-ap.ansell.com) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, February 24, 2003 at 03:14:57 ---------------------------------------------------------------------------
Email: pgan-at-ap.ansell.com Name: Gan Phay Fang
Organization: Ansell Shah Alam Sdn Bhd
Education: Graduate College
Location: Shah Alam,Selangor, Malaysia
Question: Dear Sir Good day ! I am a beginer as a SEM user. Currently, I notice that the higher the magnification of the SEM , the lesser the penetration of the electron beam as shown by the EDAX spectrum. It would be nice if you could tell me whether there is any correlation between the EDAX and the magnification as well as between the EDAX and the sharpness of the SEM imej.
Below is the result of your feedback form (NJZFM-ultra-55). It was submitted by (dha6n-at-virginia.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, February 24, 2003 at 13:27:55 ---------------------------------------------------------------------------
Email: dha6n-at-virginia.edu Name: Dalaver Anjum
Organization: University of Virginia
Education: Graduate College
Location: charlottesville, VA 22904
Question: I'm interested to know the lattest software packages for nano-diffraction in TEM particularly for LACBED/CBED simulations. Will you please provide me some names of these? Thank you very much.
Hello, Has anyone been able to calibrate an ultrasonic bath successfully as described in the ASTM Method D5755-02? If so, what brand of sonicator are you using? Pronda Few
There should be no difference in the depth of penetration as a function of magnification. It will vary according to beam voltage (or effective beam voltage).
I am curious why you think there would be a difference in penetration depth as a function of magnification. What evidence did you see for it? I would guess that you are seeing an underlying layer disappear as you go to higher magnifications.
I would suspect that you might be getting increased charging as you got to higher magnifications and pump the same current into a smaller area. If so, that will reduce the effective beam voltage and you would get less penetration. You should also be getting artifacts in your image. I suggest looking at the high energy limit of your spectra taken at the various magnifications. The background should tail off at the energy of your beam. But say you were using a 10 kV beam but your background tailed off at 8 kV, then your effective beam voltage is only 8 kV because your sample has charged up to 2000 V.
Check it out and let us know what is happening.
Warren
At 02:38 PM 2/24/03 -0600, you wrote: } Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (pgan-at-ap.ansell.com) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, February } 24, 2003 at 03:14:57 } --------------------------------------------------------------------------- } } Email: pgan-at-ap.ansell.com } Name: Gan Phay Fang } } Organization: Ansell Shah Alam Sdn Bhd } } Education: Graduate College } } Location: Shah Alam,Selangor, Malaysia } } Question: Dear Sir } Good day ! I am a beginer as a SEM user. Currently, I notice that the } higher the magnification of the SEM , the lesser the penetration of the } electron beam as shown by the EDAX spectrum. It would be nice if you could } tell me whether there is any correlation between the EDAX and the } magnification as well as between the EDAX and the sharpness of the SEM imej. } } Thanks.
------------------------------------------- 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
It is the magnetic field from the deflection coils in the CRT monitor. Either get a shielded (more costly) computer monitor or an LCD flat panel.
gary g.
At 09:00 AM 2/24/2003, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hello Randy, I have stained thick epoxy (Polybed 812) sections (1-5 microns) annealed to glass slides with Richardson's Stain (a toluidine blue look-alike) and then re-embedded the sections for thin sectioning and observed no problems as a result of staining. Of course, I stained the sections not the tissue. I am curious as to what you hear from other microscopists. I have worked with non-osmicated tissue and it is difficult to work with when you can't see it! Dean Abel Biological Sciences 141 BB University of Iowa Iowa City IA 52242-1324
At 01:19 PM 2/24/2003 -0600, you wrote:
} Dear Listers, } We have a client looking at individual muscle fibers which need } to be fixed and embedded for immunolabeling and which will require } longitudinal sections along at least of portion of their miniscule } lengths. These things are so tiny that they are barely visible with the } naked eye in the fixative. They almost look like little dust motes, if } we can see them at all. Immuno means no osmium, so the problem arises } about how we find them once they're in the resin (probably LR White), let } alone position them precisely enough to do long sections. } Ideas we're considering include: 1) tying the fibers along a tiny } section of dark thread so we can at least see the thread; 2) staining } them with an LM stain, such as toluidine blue, before embedding them; and } 3) settling them down onto poly-l-lysine coated Thermonox cover slips to } stabilize them, then scribing the coverslips and/or using an LM } stain. We are trying to avoid pre-embedding labeling at the client's } request, but that will be an option if all else fails. Then we can } osmicate following the labeling process. } Questions: Has anyone ever tried adhering muscle fibers to poly } lysine coverslips? If so, is the attachment stable enough to get through } the processing/embedding process? Is the use of an LM stain practical, or } does this introduce a lot of contamination at the EM level? Does } toluidine blue penetrate past the fiber membrane? (I've never looked at } LM stained materials in a TEM before.) We're trying to get an idea of } what might work before we start, since there is a substantial amount of } time-consuming dissection, etc., involved. Thanks much for any ideas and } thoughts. } Randy Tindall } EM Specialist } Electron Microscopy Core---We're the Fun Core! } W122 Veterinary Medicine } University of Missouri } Columbia, MO 65211
Hi Randy, Try 1%tannic acid, it preserves antigenicity, good for membranes and imparts a pale hue to locate, embed and orient specimen easily. I haven't tried on muscle fibers though, should work.
shashi
The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear Listers,
We have a client looking at individual muscle fibers which need to be fixed and embedded for immunolabeling and which will require longitudinal sections along at least of portion of their miniscule lengths. These things are so tiny that they are barely visible with the naked eye in the fixative. They almost look like little dust motes, if we can see them at all.
Immuno means no osmium, so the problem arises about how we find them once they're in the resin (probably LR White), let alone position them precisely enough to do long sections.
Ideas we're considering include: 1) tying the fibers along a tiny section of dark thread so we can at least see the thread; 2) staining them with an LM stain, such as toluidine blue, before embedding them; and 3) settling them down onto poly-l-lysine coated Thermonox cover slips to stabilize them, then scribing the coverslips and/or using an LM stain.
We are trying to avoid pre-embedding labeling at the client's request, but that will be an option if all else fails. Then we can osmicate following the labeling process.
Questions: Has anyone ever tried adhering muscle fibers to poly lysine coverslips? If so, is the attachment stable enough to get through the processing/embedding process? Is the use of an LM stain practical, or does this introduce a lot of contamination at the EM level? Does toluidine blue penetrate past the fiber membrane? (I've never looked at LM stained materials in a TEM before.)
We're trying to get an idea of what might work before we start, since there is a substantial amount of time-consuming dissection, etc., involved.
Thanks much for any ideas and thoughts.
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We're the Fun Core! W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414
===== Shashi Singh Scientist Centre for Cellular and Molecular Biology Hyderabad-500 007 INDIA PH-91-40-7192575,7192761,7192615 FAX-91-40-7160591, 7160311
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Anyone remember Tips & Tricks... Anyway, there are a couple of discussions for locating "invisible" samples in the TEM section at the following url:
http://www.biotech.ufl.edu/EM/tips/tem.html
Good luck
Scott Whittaker Laboratories of Analytical Biology Smithsonian Institution National Museum of Natural History PO Box 37012 MRC104 Washington DC 20013-7012 202-357-1651
} } } "Tindall, Randy D." {TindallR-at-missouri.edu} 02/24/03 02:19PM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear Listers,
We have a client looking at individual muscle fibers which need to be fixed and embedded for immunolabeling and which will require longitudinal sections along at least of portion of their miniscule lengths. These things are so tiny that they are barely visible with the naked eye in the fixative. They almost look like little dust motes, if we can see them at all.
Immuno means no osmium, so the problem arises about how we find them once they're in the resin (probably LR White), let alone position them precisely enough to do long sections.
Ideas we're considering include: 1) tying the fibers along a tiny section of dark thread so we can at least see the thread; 2) staining them with an LM stain, such as toluidine blue, before embedding them; and 3) settling them down onto poly-l-lysine coated Thermonox cover slips to stabilize them, then scribing the coverslips and/or using an LM stain.
We are trying to avoid pre-embedding labeling at the client's request, but that will be an option if all else fails. Then we can osmicate following the labeling process.
Questions: Has anyone ever tried adhering muscle fibers to poly lysine coverslips? If so, is the attachment stable enough to get through the processing/embedding process? Is the use of an LM stain practical, or does this introduce a lot of contamination at the EM level? Does toluidine blue penetrate past the fiber membrane? (I've never looked at LM stained materials in a TEM before.)
We're trying to get an idea of what might work before we start, since there is a substantial amount of time-consuming dissection, etc., involved.
Thanks much for any ideas and thoughts.
Randy
Randy Tindall EM Specialist Electron Microscopy Core---We're the Fun Core! 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/
I agree entirely with Warren. Depth of penetration is independent of magnification. If you are working with a sample that you believe is stoichiometrically homogeneous in the x,y and z axes, then you should not see any difference in spectra. However, if your elemental spacing is such that you do not scan across a region that has one of the elements you are looking for, you won't see it and you may simply not be scanning a large enough area. This is particularly important in quantification. A good example of this is metal alloys wherein you may get regions with only one element in it at high magnifications. Hope that's clear.
Maybe if you state what your sample is the problem will become more clear to everyone.
Regards, Peter Tomic
-----Original Message----- } From: Warren E Straszheim [mailto:wesaia-at-iastate.edu] Sent: Monday, February 24, 2003 4:30 PM To: Microscopy-at-sparc5.microscopy.com Cc: pgan-at-ap.ansell.com
There should be no difference in the depth of penetration as a function of magnification. It will vary according to beam voltage (or effective beam voltage).
I am curious why you think there would be a difference in penetration depth as a function of magnification. What evidence did you see for it? I would guess that you are seeing an underlying layer disappear as you go to higher magnifications.
I would suspect that you might be getting increased charging as you got to higher magnifications and pump the same current into a smaller area. If so, that will reduce the effective beam voltage and you would get less penetration. You should also be getting artifacts in your image. I suggest looking at the high energy limit of your spectra taken at the various magnifications. The background should tail off at the energy of your beam. But say you were using a 10 kV beam but your background tailed off at 8 kV, then your effective beam voltage is only 8 kV because your sample has charged up to 2000 V.
Check it out and let us know what is happening.
Warren
At 02:38 PM 2/24/03 -0600, you wrote: } Below is the result of your feedback form (NJZFM-ultra-55). It was } submitted by (pgan-at-ap.ansell.com) from } http://www.msa.microscopy.com/Ask-A-Microscopist.html on Monday, February } 24, 2003 at 03:14:57 } --------------------------------------------------------------------------- } } Email: pgan-at-ap.ansell.com } Name: Gan Phay Fang } } Organization: Ansell Shah Alam Sdn Bhd } } Education: Graduate College } } Location: Shah Alam,Selangor, Malaysia } } Question: Dear Sir } Good day ! I am a beginer as a SEM user. Currently, I notice that the } higher the magnification of the SEM , the lesser the penetration of the } electron beam as shown by the EDAX spectrum. It would be nice if you could } tell me whether there is any correlation between the EDAX and the } magnification as well as between the EDAX and the sharpness of the SEM imej. } } Thanks.
------------------------------------------- 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
Randy, We have used the wire loop and formvar film trick for arabidopsis roots that are also extremely small and impossible to see.
Make some small loops out of very thin copper wire leaving a "handle" for future manipulation. Cast a formvar film on a glass slide as is normally done but cut it into squares prior to floating the film off of the slide. Pick up the film squares with the wire loops so that you have a coated loop.
Next lay your fixed muscle fiber on to the film-covered loop. It should adhere fairly nicely.
Final step is to again sandwich the fiber with a formvar film. This takes a bit of practice as you don't want to dislodge your fiber. Just come down from above with your loop and the fiber clinging to the lower surface so it hits the new formvar film piece rather than the water.
The loops + film + fiber can then be carried through all the remaining solutions and even embedded in a flat bottomed capsule. The wire can be dug out o fthe polymerized resin leaving the fiber. The remaining block can then be cut off and reoriented if necessary.
Care does have to be taken so as to keep the film intact. It is sometimes helpful to stick the "handle" down into some wax melted into the bottom of a jar and then gently add and subtract solutions from the jar. In this way you are sure that the formvar film will not touch anything. Since it is a double or triple layer (we sometimes use a rectangular piece of formvar film then applying the final cover so that it actually covers both sides of the wire) it will withstand the s;urface tension changes as you rasie and lower fluid volumes.
Try it...it really works great for very small hard to see tissue.
By the way, I have also used toluidine blue to pre-stain tissue prior to embedding. It seems to work reasonably well without interfering later on although you sometimes loose a fair amount of the stain when dehydrating and infiltrating.
Debby
Debby Sherman, Manager Phone: 765-494-6666 Life Science Microscopy Facility FAX: 765-494-5896 Purdue University E-mail: dsherman-at-purdue.edu S-052 Whistler Building 170 S. University Street West Lafayette, IN 47907
On 2/24/03 8:02 PM, "Dean Abel" {dean-abel-at-uiowa.edu} wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Hello Randy, } I have stained thick epoxy (Polybed 812) sections (1-5 microns) } annealed to glass slides with Richardson's Stain (a toluidine blue } look-alike) and then re-embedded the sections for thin sectioning and } observed no problems as a result of staining. Of course, I stained the } sections not the tissue. I am curious as to what you hear from other } microscopists. I have worked with non-osmicated tissue and it is difficult } to work with when you can't see it! } Dean Abel } Biological Sciences 141 BB } University of Iowa } Iowa City IA 52242-1324 } } At 01:19 PM 2/24/2003 -0600, you wrote: } } } Dear Listers, } } We have a client looking at individual muscle fibers which need } } to be fixed and embedded for immunolabeling and which will require } } longitudinal sections along at least of portion of their miniscule } } lengths. These things are so tiny that they are barely visible with the } } naked eye in the fixative. They almost look like little dust motes, if } } we can see them at all. Immuno means no osmium, so the problem arises } } about how we find them once they're in the resin (probably LR White), let } } alone position them precisely enough to do long sections. } } Ideas we're considering include: 1) tying the fibers along a tiny } } section of dark thread so we can at least see the thread; 2) staining } } them with an LM stain, such as toluidine blue, before embedding them; and } } 3) settling them down onto poly-l-lysine coated Thermonox cover slips to } } stabilize them, then scribing the coverslips and/or using an LM } } stain. We are trying to avoid pre-embedding labeling at the client's } } request, but that will be an option if all else fails. Then we can } } osmicate following the labeling process. } } Questions: Has anyone ever tried adhering muscle fibers to poly } } lysine coverslips? If so, is the attachment stable enough to get through } } the processing/embedding process? Is the use of an LM stain practical, or } } does this introduce a lot of contamination at the EM level? Does } } toluidine blue penetrate past the fiber membrane? (I've never looked at } } LM stained materials in a TEM before.) We're trying to get an idea of } } what might work before we start, since there is a substantial amount of } } time-consuming dissection, etc., involved. Thanks much for any ideas and } } thoughts. } } Randy Tindall } } EM Specialist } } Electron Microscopy Core---We're the Fun Core! } } W122 Veterinary Medicine } } University of Missouri } } Columbia, MO 65211 } } }
} Email: dha6n-at-virginia.edu } Name: Dalaver Anjum } } Organization: University of Virginia } } Education: Graduate College } } Location: charlottesville, VA 22904 } } Question: I'm interested to know the lattest software packages for } nano-diffraction in TEM particularly for LACBED/CBED simulations. Will you } please provide me some names of these? Thank you very much.
I favor the plane-wave multislice implementation by Earl Kirkland. In the frozen phonon approximation, it is arguably the most accurate algorithm (see D. A. Muller et al Ultramicroscopy 86, 371 (2001)). Best of all, you get the entire source code and executables for Mac and Windows for the price of Kirkland's book! The book is "Advanced Computing in Electron Microscopy", by Earl J. Kirkland, Plenum 1998, ISBN 0-306-45936-1. It contains a concise summary of electron scattering and image formation, an extensive treatment of the plane-wave multislice image simulation method, and advice for doing accurate simulations with examples.
This package has one drawback for some users: the user-interface is command-line only. There is no slick GUI and no graphical help constructing atomic models.
Good luck! Paul Voyles
Assistant Professor Materials Science and Engineering Department University of Wisconsin - Madison 1509 University Ave. Madison, WI 53706-1595 Voice: (608) 265-6740 Fax: (608) 262-8353 voyles-at-engr.wisc.edu www.engr.wisc.edu/mse/faculty/voyles_paul.html
I would like to hear from anyone with experience in using digital imaging plates in TEM and how you think this compares to using digital camera systems. Thanks.
Tom Bargar Electron Microscopy Core Research Facility 986395 Nebraska Medical Center Omaha, NE 68198-6395
The 8th Annual RMC Materials Microtomy Course & Workshop is hosted by Boeckeler Instruments in Tucson, Arizona from April 29 - May 2, 2003. Designed specifically for materials scientists needing exposure to advances in specimen preparation for electron microscopy, this is a "hands-on" course catering to all levels of experience. Full details can be seen at www.rmcproducts.com
I have a question for those doing live fluorescent cell imaging in a controlled environment over a period of many hours:
We have a plexiglass incubator installed around the stage of a Nikon inverted microscope, setup for live cell imaging and are looking for a device to both regulate and measure the amount of CO2 inside the chamber. Does anyone know of such a product? Phenol red in culture media can interfere with optical quality, yet we have to maintain a stable pH in the solution. I prefer not to have a probe directly in the dish because often the dish will be closed, therefore, measuring the atmosphere is more practical.
Also, thinking about which gas to use, I thought of a pure CO2 tank but someone said it may make the environment hypoxic if the exhaust is too close to the dish. If the end of the tube (CO2 is bubbled through water) is too distant from the dish, the gas can escape through numerous gaps in the setup and we'll be going through tanks on a daily basis. Would a 5% CO2/air mixture be better?
Thanks for any help Judy
Judy Trogadis Bio-Imaging Coordinator St. Michael's Hospital, 8Queen 30 Bond St. Toronto, ON M5B 1W8 Canada ph: 416-864-6060 x6337 pager: 416-685-9219 fax: 416-864-6043 trogadisj-at-smh.toronto.on.ca
Dear Tom It was huge discussion on ListSerrver about this issue a year or so ago. You may probably find that discussion in the archive. My personal opinion, which I explained in past discussion and may explain again is briefly that image plates are superior in terms of linearity and sensitivity (not necessary better than modern digital cameras). You may load them in the standard film holder and share between the instruments (impossible for digital cameras). The downside of the plates: you need to read them relatively quick after exposure. Scanning will take about 2 min per plate and you have to upload plates from the film holders and load into the scanner and then load them back to film-holder for re-using. So, it's a lot of technical work loading-uploading-scanning etc. I am not sure but it seems to me you have to load-upload in the dark (may be not). In general, image plates may deliver more pixels than moderate digital cameras.
Digital cameras are attached to the instrument, you could not share them. From another hand it always ready: you don't need to load-upload-scan etc as it happening with image plates (if somebody before you uses all plates in the scope for instance). Most moderns digital cameras gives you the chance to keep your film in the scope as well, so you may use film or digital camera without any changes and camera chamber ventilation/vacuuming. The down side of most digital cameras that they do provide less pixels than image plate (not all of them, top models has similar amount of pixels like Ultrascan 4000 from Gatan). Another advantage which appeared to me only when I start using the digital camera on my own is that vacuum in the scope becomes better (there is no frequent ventilation/vacuuming of the camera chamber). Another beauty of the digital cameras (yes, I am voting for digital cameras) is that you have immediate access to the image - you could take the picture and immediately sent it to the printer or to the collaborator. Moreover, you may use Internet and create "video-conference" when people on the opposite side of globe will see exact the same on their screens that you see with digital camera in your microscope room. Personally, I do find that ability to see "live" image on the screen has a great educational potential. It's much easier to teach students how to focus using live image on the screen. I also use that "video-concerning" feature to work with my collaborators on frequent base. Actually, the image from digital camera permanently transmitted to our local network and everyone could see what happening in the microscope.
Personally, I am happy owner of the Gatan's BioScan 600W top mount camera. It's great camera for biological applications. I don't have any commercial interest in Gatan company, just very satisfied user.
Feel free to contact me if you have some further questions. And the very last: do not make final decision unless you will see the stuff in work. Ask for demo, took pictures and compare side-by-side. If company refuse to make demo to you - it looks suspicious to me, I would avoid such company. You may also ask company for references and how many instruments is working in your area. Another things to consider is quality of customer service and avliability (?spell) of it (how many engineers perform service in your area). Godd look.
Sergey
At 08:39 AM 2/25/2003, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry Box 951737 Los Angeles, CA 90095-1737
Has anyone tried compiling the source code from Kirkland's book under Linux? The book says it is standard ANSI C, but I get a number of errors under Caldera OpenLinux Workstation 3.1.1 with gcc 2.95 using the -ansi switch. Primarily these refer to missing trigonometric functions. Reading the documentation for gcc did not help but a look at the include directories revealed a file "tgmath.h" which has the trig-math declarations. Adding #include {tgmath.h} to the source code removed some of the compiler errors, but I still get number of errors relating to sqrtl and other functions esp. from slicelib.c.
If someone has already solved this problem I would like to know the compiler options and changes required to the source code. I do understand that my Linux and gcc are a bit dated. Has it worked directly under a recent version like SuSE Linux 8.1 with gcc 3.0, which is what I am planning to upgrade to?
Thanks for any responses, Divakar
---- Dr R Divakar Physical Metallurgy Section MCG-IGCAR, Kalpakkam 603102, India ----
-----Original Message----- } From: Paul Voyles [SMTP:voyles-at-engr.wisc.edu] Sent: Wednesday, February 26, 2003 10:14 AM To: Microscopy-at-sparc5.microscopy.com Cc: dha6n-at-virginia.edu
} Email: dha6n-at-virginia.edu } Name: Dalaver Anjum } } Organization: University of Virginia } } Education: Graduate College } } Location: charlottesville, VA 2