Hello, there, Does anyone out here know that can we put alkylamine to the turbo pump when carbon supporting film is being glow discharge? Thanks.
Peiyi
Krebs Institute for Biomolecular Research University of Sheffield Firth Court Western Bank Sheffield Yorkshire S10 2UH United Kingdom Tel: +44 (0)114 222 2000 Direct: +44 (0)114 222 2739 FAX: +44 (0)114 272 8697 E-mail: p.wang-at-sheffield.ac.uk
Briget wrote: =========================================================== I will like to do cryomicrotomy of PP wires. My question is : can anyone suggest what kind of epoxy is appropriate for cryomicrotomy? I used Epoxy Mount and it was chattered during cryo process. The manufacturer confirm to me that epoxy mount is not appropriate for cryomicrotomy. Can anynone help me? =========================================================== I am assuming you mean polypropylene coated wires and not polypropylene monofilament. We have generally found that for coated wires samples, we like to Pt coat it first (by sputtering), then embed in SPI-Pon™ 812 resin. I would expect that any of the other "Epon® 812 substitutes", available from the other major EM supplies firms would work just as well.
You will definitely want to use a diamond knife on this and you can vary the hardness of the resin in way that gives you the best sections. Use a knife angle that is not larger than 45°, the lower the temperature (usually) the better.
Disclaimer: SPI Supplies offers for sale the resin and diamond knives mentioned and performs this kind of cryoultramicrotomy for clients.
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.spi.cc ######################## ===========================================
Hamamatsu recently introduced a new camera called the electron bombardment ccd where accelerated electrons directly bombard a back thinned, peltier cooled ccd. The electrons are emitted from a photocathode for applications particularly in low light microscopy. With a full well capacity of 300,000 electrons, I wonder if this approach could be used in direct exposure of the ccd to the electron beam. Shane
K. Shane Collins Scientific Instrument Company 805.444.4953 cell 310.568.9188 office 310.568.9189 fax
-----Original Message----- } From: Paul Voyles [mailto:voyles-at-research.nj.nec.com] Sent: Friday, May 26, 2000 9:11 AM To: Microscopy-at-sparc5.microscopy.com
} You could probably expose your CCD chip directly to the electron bean -- } and buy a new chip every few hundred exposures or so! Electrons have
There is an even worse problem with exposing the CCD directly the electron beam. The p-n junctions in the CCD chip have a certain "well capacity" - a number of electron/hole pairs they can hold before they saturate. Fast (keV) electrons are much more efficient at producing electron/hole pairs than photons - so much so that a pixel on the CCD would saturate after about 15 fast electrons hit it. Just the square root N shot noise at that level is about 25% - larger than typical TEM micrograph contrast of 10-20%. That's why a phosphor or scintillator is necessary to transform the fast electrons into photons.
Yes, it could be done "in theory". Somebody would need to figure out the software and perhaps modify the hardware. Then we would find that the total exposure of the specimen to the electron beam maybe a muliple of the film's exposure. Afterall, an 8 sec film exposure would not amount in digital to 10x0.8, but we would require considerable time in between exposures. Since the problems in the discussed circumstances are specimen movement and beam damage, it seems that taking multiple exposures is a poor option.
Digital cameras are for some situation too sensitive to electron exposure. Cutting back on electrons is no option since its the electrons that form the image in the first instance. Much easier in light microscopy . . . insert a neutral density filter. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
} Hello Jim, } } just a couple of remarks to your email. } } your remark 1) What you can do of course with digital imaging (provided } you have the necessary hardware), is to automatically collect larger } areas by taking several images that overlap, reducing the advantage that } film has in this area and perhaps offer other possibilities as I just } explained in another posting as a response to Bill Tivol's posting. } } your remark 2) I am not sure you are not comparing apples and oranges. } What you are saying is, that because of the "slowness" of film you need } longer exposures, thereby averaging out the statistical noise of the } electron, which is not the case for CCD cameras at short exposures, } hence they appear more noisy. In essence what you are doing is to } compare a short exposure image to a long exposure image. What you can do } with a CCD camera is the following: You can get (perhaps) real-time dark } field images and position your sample and/or decide if you want to } actually take the image. Then you take a SERIES of images, let's say 10, } each at an exposure time of 1/10 of the film exposure. This can be done } automatically, of course. Finally, you add or average all of these } images using a pattern recognition to align them first. The result: A } dark field image that should be as noisy as the film image, with much } less problems due to drift during long exposures, a higher dynamic range } and visible immediately on the viewing screen. } } Michael } } Michael Bode, Ph.D. } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } =================================== } phone: (888) FIND SIS } (303) 234-9270 } fax: (303) 234-9271 } email: mailto:info-at-soft-imaging.com } web: http://www.soft-imaging.com } =================================== } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } Sent: Tuesday, May 30, 2000 7:13 PM } To: 'William Tivol' } Cc: microscopy-at-sparc5.microscopy.com } Subject: RE: Film vs Digital } } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } Peter Bond is right in saying "The future is with digital image capture" } if we } were at some future date to count heads of digital versus film TEM } users. } That change will be for reasons of convenience and to save labour. These } are } powerful and valid reasons. } } Bill Tivol has given one set of applications where digital currently } does not } always measure up to film. } Here are another couple of such applications. } 1 When great enlargements are required film is superior. This is } because of } film's greater resolution, but more importantly, because its much, much } easier } to take images at moderate powers and highly enlarge. That way we take } advantage of the TEM's greater depths of focus at low powers and of } film's } higher resolution/ image detail. } 2 Whenever a TEM image is taken at low brightness (to avoid beam } damage or at } very high powers or in dark-field) relatively few electrons form the } image and } make that image grainy. Film is very slow and requires then a longer } exposure, } thus boosting the quantity of electrons used and improving the image. } Digital } is much more sensitive and so the exposure must be shortened. As a } result the } best digital camera will record, quiet faithfully the grainy, unsharp } image. } } Many labs rarely or never use such applications and for them the reasons } to } change to digital now may be overwhelming. } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com } } On Wednesday, May 31, 2000 5:18 AM, William Tivol } [SMTP:tivol-at-wadsworth.org] } 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 } } } -----------------------------------------------------------------------. } } } } } } Peter Bond wrote: } } } } } The continuing discussion on whether film is better than digital, } and } } } whether we can directly compare their resolving ability is very } interesting } } } and throwing up some really useful technical stuff. But aren't we } missing } } } the point a bit? Surely the image quality is user defined - if you } or your } } } customer, are satisfied with the end product then the equipment has } done } } } its job. How often does one need to examine a micrograph to assess } its } } } limits? If are getting that close to a picture then you may be } taking } } } things out of context a bit and losing the whole concept. } } } } } } The future is with digital image capture, let's hope the price of } the } } } equipment comes down to something more attainable! } } } } } } } Dear Pete, } } If one is doing quantitative image processing, the better } } resolution } } available from film can be relevant. For example, if one wants to do } } corellation averaging, one needs as many objects in the picture as } possible, } } and also as good resolution as needed--e.g., for 1 nm resolution of } the } } reconstruction, a pixel size of 0.25 nm times the magnification is } necessary, } } so the recording medium must have resolution equal to or better than } that. } } The pixel size of the scanner must also be this small--obviously } irrelevant } } for digital recording. A 5 micrometer scanning-pixel size at 20kx mag } } gives a pixel resolution of 0.25 nm, and for 6.5 by 9 cm film, one } will have } } about 12,000 by 16,000 (taking the header into account) pixels of } useful } } information. This will give a broader area than that available at } equal } } resolution from any CCD chip now out there. I can assure you that I } have } } often had to determine the information limits in a particular } micrograph. } } Yours, } } Bill Tivol } } } } }
The electron energies used in this "intensifier" will be in the order of one or several kV. In TEM however the energies are much higher thus one incident electron will generate for example hundred electron-hole pairs in the CCD thus saturating it very fast. Another factors are damage to the CCD chip and X-Rays.
I was thinking about another approach. A chip consisting of matrix of thermo-sensitive elements. Above each element there will be a metal block with height equal or larger than the stop path for the electron energy used. The whole thing will be cooled in a similar way as the CCDs. When this assembly is exposed to the beam each block will increase its temperature depending on the number of electrons stopped. After the exposure the matrix is scanned and the temperature increase at each element is measured (ofcourse before each exposure a reference image has to be taken).
The benefits: - Very high efficiency. Almost every incident electron will contribute to the image. - Huge dynamic range. - Linearity (after the temperature-signal characteristic of each element has been calibrated) - Narrow point spread function (maybe).
Problems: - Difficult to manufacture (the metal blocks should be insulated from each other) - Maybe low sensitivity. I haven't calculated how much the temperature increase will be (for example 5x5x30um Cu block hit by one 300 kV electron) but I suspect it will be very small. Also It will depend on the sensitivity of the thermo-measuring elements. - Saturation. After each exposure one has to wait some time for the thing to cool down again.
There are maybe other difficulties which do not come in mind now.
Hmmm now I start thinking about X-Rays. Actually the main part of the incident energy will go into X-Rays thus making this device useless.
Rado
--------------------------------------------------------------------- Radostin Danev Laboratory of Ultrastructure Research National Institute for Physiological Sciences Myodaiji-cho, Okazaki 444-8585, JAPAN e-mail: rado-at-nips.ac.jp --------------------------------------------------------------------- ----- Original Message ----- } From: Shane Collins {kshanec-at-gte.net} To: Paul Voyles {voyles-at-research.nj.nec.com} ; {Microscopy-at-sparc5.microscopy.com} Sent: Thursday, June 01, 2000 9:15 AM
AnalySIS has a module which does this see Soft Imaging's web site at http://www.soft-imaging.de Chris
To: Microscopy-at-sparc5.microscopy.com } From: Jim Ferreira {ferreira1-at-llnl.gov}
===================================================================== DR CHRIS JEFFREE BIOSEM - BIOLOGICAL SCIENCES EM FACILITY UNIVERSITY OF EDINBURGH Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 131 650 5345 FAX. #44 131 650 6563 Mobile 0410 585 401 email c.jeffree-at-ed.ac.uk SEM / TEM bookings sem-at-ed.ac.uk =====================================================================
In a message dated 5/31/00 2:52:24 PM, ferreira1-at-llnl.gov writes:
Anyone aware of software capable of generating depth profiles from digital SEM stereo pair images? I am aware of the Oxford ISIS system/program, but wondered if other companies offer similar software that would run 'stand-alone' in a PC and work with any digital files.
Jim I have been using uMex software for the last 6 months which does exactly what you want. Given a stereo pair it will produce a 3D surface reconstruction. It has a function that enables images to the two images to be manually aligned prior to the calculation. This function when used correctly seems to result in faster calculations.
Another other nice feature is that images can be output in VRML file format. So you can view the images with shareware 3D packages. We use a SGI for viewing as its rendering speed is significantly faster than a PC.
I suggest you check out the following web site. http://www.alicona.com/en/products.htm
Regards Colin MacRae
************************************************************************ Manager of Electron Microscopy Group (Clayton)
I've been following the thread on film vs digital. We all know that the resolving power of digital CCD faceplates is approaching that of conventional film emulsion, but isn't equivalent yet. Can someone remind me of the number of mega-pixels that a CCD will need to equate to ISO 100 print film, ISO 64 or ISO100 slide film and the highest resolving B&W film of all, Technical Pan at ISO 25 and ISO 100? If anyone out there can lead me through the logic and the maths, I'm sure others will also find it helpful. I'll repost a summary of the replies that I get.
Regards, Jeremy Sanderson
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} Hamamatsu recently introduced a new camera called the electron bombardment } ccd where accelerated electrons directly bombard a back thinned, peltier } cooled ccd. The electrons are emitted from a photocathode for applications } particularly in low light microscopy. With a full well capacity of 300,000 } electrons, I wonder if this approach could be used in direct exposure of the } ccd to the electron beam. } Shane
This certainly could be a significant improvement in CCD imaging. Most of the width of the point spread function of current CCD camera is due to photon spread in the scintillator, so presumably removing the scintillator would allow digital images at resolutions very close to the pixel size of the CCD chip.
I'm not familiar with the Hamamatsu camera, but I know that the Gatan camera I currently use has a CCD with a full well capacity of ~500,000 electrons. In order for the Hamamatsu chip to work they would have to find some way to reduce the electron/hole yield of the incident fast electrons - maybe with a very thin CCD chip?
Paul Voyles
} K. Shane Collins } Scientific Instrument Company } 805.444.4953 cell } 310.568.9188 office } 310.568.9189 fax } } } -----Original Message----- } From: Paul Voyles [mailto:voyles-at-research.nj.nec.com] } Sent: Friday, May 26, 2000 9:11 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: RE: new developments in imaging systems? } } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} Dear All, } } I've been following the thread on film vs digital. } We all know that the resolving power of digital CCD } faceplates is approaching that of conventional film } emulsion, but isn't equivalent yet. } Can someone remind me of the number of mega-pixels } that a CCD will need to equate to ISO 100 print film, } ISO 64 or ISO100 slide film and the highest resolving } B&W film of all, Technical Pan at ISO 25 and ISO 100? } If anyone out there can lead me through the logic and } the maths, I'm sure others will also find it helpful. } I'll repost a summary of the replies that I get. } } Regards, Jeremy Sanderson }
About twenty-five million pixels in a 35mm Kodachrome slide is the number I have heard from several sources, none of which I can remember now.
Geoff -- ********************************************** 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 **********************************************
No, it's not really a problem. It's been done with low density microscopy all the time. Granted, there are some technical aspects to be overcome, but (and I can only speak for ourselves) we have done that on a number of microscopes. You are of course correct, that 10 images at 0.8 seconds take longer than 8 seconds as the image has to be transferred, etc. BUT: that's what beam blankers are for. It is pretty straightforward to take an image at 0.8 seconds, then blank the beam very quickly before taking the next image. That way you get pretty close to the 8 sec total exposure. If there is no beam blanker on the microscope, in most cases it can be added.
I am not sure what you mean by "too sensitive". The cameras are usually constructed so that 1 electron from the beam creates between a few tenth to a few counts (these are all statistical data, of course). The well width divided by this sensitivity then determines, how many primary electrons are needed to fully expose one pixel. For example, if the well width is 50,000 electrons and the sensitivity is 1 count/electron, one needs 50,000 primary electrons to fill the well. This translates into roughly a 0.4% statistical error.
} From a practical standpoint: You can take images with most cameras when the exposure meter on the microscope reads a couple of seconds without overexposing the camera. On the other hand, you can reduce the intensity of the beam until you see single electron events.
The one area where CCD cameras may be too sensitive is diffraction. The normally huge intensity in the transmitted beam often leads to saturation. In CCDs this can lead to blooming (the intensity spills over into neighboring pixels). This can be taken care of with special chips that have anti-blooming features, but this usually has some other drawbacks. Again, this can also be overcome somewhat with multiple exposures. Film behaves more civilized here, as it simply stops responding to the electrons, but this makes film more or less useless for quantitative measurements of diffraction patterns. I have done diffraction with CCDs many times and though it does require some tweaking, one can get very good results from them.
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: jim [mailto:jim-at-proscitech.com.au] Sent: Wednesday, May 31, 2000 9:37 PM To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com'
Yes, it could be done "in theory". Somebody would need to figure out the
software and perhaps modify the hardware. Then we would find that the total exposure of the specimen to the electron beam maybe a muliple of the film's exposure. Afterall, an 8 sec film exposure would not amount in digital to 10x0.8, but we would require considerable time in between exposures. Since the problems in the discussed circumstances are specimen movement and beam damage, it seems that taking multiple exposures is a poor option.
Digital cameras are for some situation too sensitive to electron exposure. Cutting back on electrons is no option since its the electrons that form the image in the first instance. Much easier in light microscopy . . . insert a neutral density filter. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
Jim- I saw an ad a while back for such software. I have since moved on to other things before I had a chance to obtain the demo software. Below is the contact information I have for the company selling the stereo image analysis software. Since this is an edited version of the message I received several months ago, the terms and conditions may have changed. If you do try this software, I, and I am sure the rest of the listserve would be very interested in hearing how well it works, as I do have an occational need for this capability.
Here is the information (edited) that I received from the company offering the software: now the evaluation version of MeX is available. The test period of MeX is limited to 6 weeks.
The evaluation version is delivered with a small database and the complete analysis tools. You can process your own images without restrictions.
We also offer to preinclude your SEM-images into the database of the evaluation version. You just have to send us an email and we give you detailed information on how you should capture your images.
If you have any questions, please do not hesitate to contact us.
Jim, you may want to check out our web site for the Stereo software. It has some images and examples of stereo evaluations from SEM images. The 'Stereo' part is not a stand-alone software, but can be combines with our analySIS Docu software for a stand-alone application. Michael
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
Hello folks Anyone aware of software capable of generating depth profiles from digital SEM stereo pair images? I am aware of the Oxford ISIS system/program, but wondered if other companies offer similar software that would run 'stand-alone' in a PC and work with any digital files. Any help or suggestions along these lines would be most appreciated. Thanks, Jim
Do not give up if you do not have an ESEM or LVSEM there may be hope yet? Do you have a SEM with a rear manifold that connects directly to the DP and do you have a BSE detector? Sorry but those SEM that have the pump directly attached to the specimen chamber are no good for this procedure. AND I must give all the credit to Viv Robinson who spawned this idea back in the 80's.
If you do have a manifold system you are in luck. Find a rubber bung that will fit into the manifold at the rear of the specimen chamber. Drill (use LN2) a 1/4" hole in the bung and then place it in the rear manifold. Switch off or better still unplug your Everhart-Thornley detector (high voltage plus poor vacuum = arcing!). Place you "wet" specimen in the microscope and pump down. The bung will spoil the vacuum in the chamber for about 20 minutes or so and imaging with the BSE detector you will have your own LV SEM.
To retain the moisture longer you may quench the specimen in LN2 before putting it into the microscope. The frost will sublime away and you will be able to watch what happens to the moisture etc.
We use this technique on all sorts of samples. A rubber bung is cheaper than buying an LVSEM and the results, if you work quickly, are pretty good.
Try it?
Steve Chapman Senior Consultant Protrain For consultancy and professional training in EM world wide Tel 44+ 1280 814774 Fax 814007 www.emcourses.com
Steve Chapman Senior Consultant Protrain For consultancy and professional training in EM world wide Tel 44+ 1280 814774 Fax 814007 www.emcourses.com
Is it just my problem, or isn't this all getting a little bit unfocussed! As far as TEM is concerned, the CCD is not directly exposed to the beam at all, so its effective resolution depends on the nature of the system that presents the image to the CCD. The two predominating technologies for transfer of the image to the CCD in TEM cameras are a fibre-optic linkagage between a phosphor or YAG scintillator and the CCD, or an optical coupling via a lens (for example the excellent f1.2 50mm Zuiko macro lens by Olympus). In both instances, the ultimate resolution of the system is probably set by the electron sensor, which is the phosphor or YAG scintillator. I suspect that fibre optic couplings probably degrade that resolution, but I say that without reference to the facts, so please correct me if I am mistaken. Optical coupling could in principle project the spatial data recorded by the phosphor or YAG to any desired magnification. It can thus be recorded by a CCD using many pixels or few depending on the optical configuration. So what do we mean by resolution in this context, when the smallest object which can be imaged by a TEM can be projected onto any desired quantity of CCD pixels?
To begin to answer Jeremy's question directly, we need to know how much detail a Technical Pan negative can record. The figures depend on processing technique and the test object luminance and contrast, but the modulation transfer function figures published by Kodak indicate that a spatial frequency in excess of 200 cycles per mm is easily recordable. For a test object with contrast 100:1 they quote 320 line pairs per mm. The CCD pixel spacing required to achieve this feat would be 640 pixels per mm. That equates to a requirement for 15360 x 23040 pixels to match the resolving power of a 24x36mm Technical Pan exposure. That's 0.35 Giga pixels in round numbers.
So what was that I heard about the death of silver imaging? I don't think so. Not for a while yet.
Best wishes Chris
} Dear All, } } I've been following the thread on film vs digital. } We all know that the resolving power of digital CCD } faceplates is approaching that of conventional film } emulsion, but isn't equivalent yet. } Can someone remind me of the number of mega-pixels } that a CCD will need to equate to ISO 100 print film, } ISO 64 or ISO100 slide film and the highest resolving } B&W film of all, Technical Pan at ISO 25 and ISO 100? } If anyone out there can lead me through the logic and } the maths, I'm sure others will also find it helpful. } I'll repost a summary of the replies that I get. } } Regards, Jeremy Sanderson } } __________________________________________________ } Do You Yahoo!? } Send instant messages & get email alerts with Yahoo! Messenger. } http://im.yahoo.com/ }
===================================================================== DR CHRIS JEFFREE BIOSEM - BIOLOGICAL SCIENCES EM FACILITY UNIVERSITY OF EDINBURGH Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 131 650 5345 FAX. #44 131 650 6563 Mobile 0410 585 401 email c.jeffree-at-ed.ac.uk SEM / TEM bookings sem-at-ed.ac.uk =====================================================================
We at Ted Pella Inc. have used our Pelco UVC2 UV Cryo Chamber to cure epoxies but most of our UV curing has been with Acrylic resins. The literature we have on Eponate Araldite does not show it to be UV curable but our chemist wouldn't be surprised if it didn't accelerate the cure. The epoxy mixture should cure overnight at 60 degrees C. We have a technical note on the use of Epon-Araldite for embedding specimens and literature on our Cryo Chamber that may help. I would be happy to fax them to you and/or have you talk with our chemist.
Mark J Armogida VP Engineering and Production Ted Pella Inc.
Tamara Howard 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 } -----------------------------------------------------------------------. } } Greetings! } } Has anyone ever cured Epon-Araldite with UV? I was given a } cobbled-together protocol by a non-microscopist post-doc; the protocol } calls for a dry-ice, UV cure of Epon-Araldite. It says it can take from } 3-7 days. I'm wondering if this is either a resin mix-up *or* the resin } just set up all by itself (over that time frame I've seen it happen), just } because it was a thin layer and left alone - nothing to do with the UV } exposure. She has no ref. for this technique and isn't sure now where she } got it. Sigh. I've only been able to find heat-cure protocols...anyone } have any leads or thoughts on this UV thing? } } (Sorry about the cross-post for those of you on both of these servers) } } Thanks! } } Tamara (Planning to use the oven.....) Howard } CSHL
I am seeking to speak to individuals who know about designing and developing applications for indentification of rare cells in microscopic biological preparations. Someone who knows how to optimize microscopy autofocusing procedures. I am more interested in speaking to an electrical engineer who is more into digital image processing. Can anyone suggest what direction to take?
We are trying to help a client find and identify a bacteriophage. His bacteria (Pasturella sp.) are plated out on agar and show well-defined clear areas with sharp edges where the phage are supposed to be. So far, we have taken carbon-coated grids and placed them gently onto the surface of the clear areas, then lifted them off and negative-stained with PTA or uranyl acetate. We have also run buffer across the clear areas, then pipetted it onto the grids and stained it. A microbiologist who works with phage in another lab has taken samples from the clear areas and concentrated them down and we have stained those also.
So far we have found exactly two phage-like organisms in a total of about 10 grids. Not a stellar performance.
We figure the possibilities are: 1) the bacteria are being killed by something other than phage; 2) we're looking for a particular type of phage that may not be there, and we're just not seeing what's actually causing the clear areas, or 3) for some reason we're just not getting the things adhering to the grids, although we've used these methods successfully many times before.
Does anyone else have any ideas that might help us out, especially on technique? Our client is almost certain that phage are present. We just can't find them.
Thanks in advance.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility 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/
One of our SEM users would like to label biofilms with lectins. Three of them. At once. I have used gold conjugated to goat anti-biotin to label biotinylated lectins for TEM in the past, so I'm hoping to follow the same kind of procedure. I have not done any immunolabeling for SEM, although our FESEM has been used for such. I would be grateful for any advice! If he wants to triple label, what sizes of gold would be useful? If he wants to quantify the three, what kinds of controls for labeling efficiency should we run? All hints ahd tips gratefully accepted!
Mahalo, 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* ****************************************************************************
I have not generally followed any discussion on gold enhancement for light microscopy (photons? I don't do photons), but now I need to ask for someone what people suggest for immunogold enhancement for confocal microscopy. We have 10nm gold left over from TEM, and it would be useful to use it for the confocal experiment. Yes, we're being cheap!
Mahalo, 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* ****************************************************************************
} From: "Chris Jeffree" {cjeffree-at-srv0.bio.ed.ac.uk} To: "Jeremy Sanderson" {jb_sanderson-at-yahoo.com} {snip} } To begin to answer Jeremy's question directly, we need to know } how much detail a Technical Pan negative can record. The figures } depend on processing technique and the test object luminance and } contrast, but the modulation transfer function figures published by } Kodak indicate that a spatial frequency in excess of 200 cycles per } mm is easily recordable. For a test object with contrast 100:1 they } quote 320 line pairs per mm. The CCD pixel spacing required to } achieve this feat would be 640 pixels per mm. That equates to a } requirement for 15360 x 23040 pixels to match the resolving power } of a 24x36mm Technical Pan exposure. That's 0.35 Giga pixels in } round numbers. } At 16 bits this is per image this is 5,662,310,400 or 5.5 gig. I don't think film is in any danger for a long time. We need at least 1 order of magnitude for storage and 2 or 3 for processing. I remember taking 4 days to process an image. And then work on the program and trigger it again.
Gordon
Gordon Couger gcouger-at-couger.com
Stillwater, OK www.couger.com/gcouger 405 624-2855 GMT -6:00
Hello all, This is the last call for a second post-doc vacancy at the University of Barcelona (Spain) to work in the frame of a TMR programme concerning UV coatings. (More details below).
Since we are expecting the Mid-Term evaluation of the project, the starting date has been delayed to next September, so we have extended the deadline for applications.
Any one interested please reply directly to paqui-at-el.ub.es and/or send applications and a CV by mail before 30 June 2000.
Kind regards
F. Peir—
************************************************************************** Laboratory: Electronic Materials and Engineering, Department of Electronics, University of Barcelona.
Duration: 12-months, starting September 2000.
Hi Jim, some comments to Your remarks within Your text:
jim schrieb:
} ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } Yes, it could be done "in theory". Somebody would need to figure out the } software and perhaps modify the hardware. Then we would find that the total } exposure of the specimen to the electron beam maybe a muliple of the film's } exposure. Afterall, an 8 sec film exposure would not amount in digital to } 10x0.8, but we would require considerable time in between exposures.
Simply choose a CCD with higher readout performance (faster), but same quality.
} Since the } problems in the discussed circumstances are specimen movement and beam damage, } it seems that taking multiple exposures is a poor option. } } Digital cameras are for some situation too sensitive to electron exposure.
Not correct at all. The only thing is to choose the correct camera for the application You work on. It is not complicated to make a digital system which collects one count per incident electron to achieve the same signal to noise as in the electron beam. This system will be less sensitive than normally sold systems but the main advantage of digital systems that You see what You get remains and You get instant results of Your work. The only problem You have to solve for these systems is to use a not very sensitive scintillator but a very high performance slow-scan CCD. So You get less visible photons from Your electron which have to be converted in one digital count. But these digital counts must be more than presently available to achieve a good statistic. Thats the reason for our new 16bit CCD (dynamic up to 65536 digits) for high performance TEM applications.
} } Cutting back on electrons is no option since its the electrons that form the } image in the first instance. } Much easier in light microscopy . . . insert a neutral density filter. } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com
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} About twenty-five million pixels in a 35mm Kodachrome slide is the } number I have heard from several sources, none of which I can remember now. } } Geoff Geoff Analysing this a little further: A 25 Mbyte image = 8.3 M pixels to cover 24x36mm or 864mm^2 this represents 98.2 pixels per mm, or 49 line pairs per mm. Pixel size 10.2 um. Resolution about 50 line pairs per mm at best.
A 25M pixels image used to capture a 24x36mm Kodachrome slide represents 28,935 pixels per mm^2 or 170 pixels per mm. This is equivalent to a maximum resolution of 85 line pairs per mm, which may be on the conservative side for Kodachrome. pixel size = 5.88um. The image will be approx. 6120x4082 pixels, generating a file size of approx 75Mb for a 24-bit (8+8+8bit) RGB image.
To record 120 line pairs per mm, which many top 35mm camera lenses can achieve, a minimum of 240 pixels per mm are required, each 4.2 um wide. This equates to 8640x5760 pixels for a 24x36mm frame = 50M-pixels or 50Mb in 8-bit greyscale, or 150Mb in 24-bit RGB.
At 320 line pairs per mm (Technical Pan) the minimum required 640 pixels per mm is a pixel size of 1.56 um
Presumably for a light image the diffraction limited resolution is approx 1/2 lambda which at 540nm is 0.27um. So looking to the future of ultimate-performance CCDs, direct recording of a diffraction limited light image projected onto the sensor requires at the very least 3703 pixels per image mm or 13,717,421 pixels per mm^2 (greyscale 8-bit)
However, if we are doing light microscopy with an NA 1.4 x100 objective, how much resolving power do we need on CCD or film?
Data is at 0.27um resolution (lambda = 540nm). Let's round this to 0.3um. Magnification at 24x36mm film image is x100, so pixels must be an absolute maximum of 30um wide to record the significant data = 33.3 pixels per mm, equivalent to 800x1200 pixels to record the whole 35mm negative area. However, most CCDs are much smaller than 35 mm frames, typically 1/3 inch. So an 800x1200 pixel CCD at 8x12 mm, 1/3 of the linear dimensions of a 35mm frame would use 9 pixels to record the smallest image details. This is about right from the point of view of resolution, but to record the whole 35mm frame we need about 2400x3600 pixels on our CCD.
Note also that Technical Pan has (depending on the criterion used to assess its performance) up to 10 times the resolving power required to record all there is to see in a diffraction-limited LM image made with a 100x NA 1.4 lens. So you can comfortably afford to use a 60x NA 1.4 lens, thereby getting the same resolution with a bigger field of view.
Many years ago, I took a photograph of a street scene using a Canon 35mm SLR loaded with Kodak Recordak (I think this was a single layer microfilm emulsion). Examined in a light microscope, the image clearly, legibly recorded the brand-name of a child's push chair. I tried to print this brand name using a DeVere point- source enlarger with an image size of 20x30 inches produced with a Schneider Componon lens, but was completely unable to produce a legible image. The point I am making here is that the combination of some high performance films, with high quality lenses of the standard produced by the leading camera manufacturers can record more detail on the film than you can easily get back out by conventional printing. I suspect the same is true of EM exposures.
So there is no contest - film beats CCDs for resolution hands down. And you can process the image on the cheap. No money goes to Intel or Microscoft, Adobe or Epson. But resolution is not primarily what we buy CCDs for. We buy them primarily for instant image capture in a format suitable for digital storage, digital transmission quantitative data recording and image processing.
Chris
===================================================================== DR CHRIS JEFFREE BIOSEM - BIOLOGICAL SCIENCES EM FACILITY UNIVERSITY OF EDINBURGH Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 131 650 5345 FAX. #44 131 650 6563 Mobile 0410 585 401 email c.jeffree-at-ed.ac.uk SEM / TEM bookings sem-at-ed.ac.uk =====================================================================
Gordon's and Chris' contribution look very bleak for digital, but the comparison is not really fair, as we should look at the practical aspects too. The unaided eye resolves lines 0.1mm apart, so to see the full (200 black+200 white) 400+ lines/mm that may be recorded on TEM film we would need to enlarge over 40x. This requires an enlarger with a very wide angle lens to print small portions of the negative and it is technically difficult to so enlarge a whole negative since a 4" negative becomes 160" or over 4m in size. That degree of enlargement is not fully useful since in a well exposed TEM negative at just under 30x electron noise becomes the problem, meaning not enough electrons contributed to the image. So enlargements beyond 30x are empty- and provide no further information. Truly not a serious problem. The practical part is that few people ever find it useful to enlarge more than 15x and most TEM images reproduced are barely the size of the original negative, however, they are enlarged from a smaller portion thereof. For these most common applications, digitals with 10+ megabytes provide excellent image details and tonal gradation. However, that size image can only cover the equivalent of a small 35mm negative equivalent and does not allow high enlargement or choosing of an adjacent field. It appears that the best of both worlds is the use of conventional TEM negatives for archiving and scanning those as required for printing.
It should be noted that this discussion concerns TEM. Digitals in SEM are less daunting and it is not problematical to produce excellent digitals, comparable with film. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Friday, June 02, 2000 11:41 AM, Gordon Couger [SMTP:gcouger-at-couger.com] wrote: } } } } From: "Chris Jeffree" {cjeffree-at-srv0.bio.ed.ac.uk} } To: "Jeremy Sanderson" {jb_sanderson-at-yahoo.com} } {snip} } } To begin to answer Jeremy's question directly, we need to know } } how much detail a Technical Pan negative can record. The figures } } depend on processing technique and the test object luminance and } } contrast, but the modulation transfer function figures published by } } Kodak indicate that a spatial frequency in excess of 200 cycles per } } mm is easily recordable. For a test object with contrast 100:1 they } } quote 320 line pairs per mm. The CCD pixel spacing required to } } achieve this feat would be 640 pixels per mm. That equates to a } } requirement for 15360 x 23040 pixels to match the resolving power } } of a 24x36mm Technical Pan exposure. That's 0.35 Giga pixels in } } round numbers. } } } At 16 bits this is per image this is 5,662,310,400 or 5.5 gig. I don't } think film is in any danger for a long time. We need at least 1 order } of magnitude for storage and 2 or 3 for processing. I remember } taking 4 days to process an image. And then work on the program and } trigger it again. } } Gordon } } Gordon Couger gcouger-at-couger.com } } Stillwater, OK www.couger.com/gcouger } 405 624-2855 GMT -6:00
The world is full of possible solutions, but are they practical.? To produce high-resolution, dark-field or any others TEM images that require more electrons to form a clear image, Mike Bode would use multiple digital exposures. The exposures could be layered and combined into one superior image. This image would be made up of more pixel and is formed by more electrons and so would be noise-free and hence could be further enlarged then otherwise possible. Perhaps. Beam blanking would largely save the specimen from beam damage and drift could be compensated for by matching up the digitals. Great. How much time is required between exposures to transfer a minimum 10mb image per exposure? What would be the total time from focusing to the last exposure? What about Z-drift in the interim requiring objective changes and what about the total cost of this additional get-up. The mind boggles at a through focus series. When pushing the limits a piece of film seems more effective, cheaper and fa ster. Again, I don't doubt that there is now a large place for digital in TEM, but its no panacea. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Friday, June 02, 2000 1:24 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] wrote: } } No, it's not really a problem. It's been done with low density } microscopy all the time. Granted, there are some technical aspects to be } overcome, but (and I can only speak for ourselves) we have done that on } a number of microscopes. You are of course correct, that 10 images at } 0.8 seconds take longer than 8 seconds as the image has to be } transferred, etc. BUT: that's what beam blankers are for. It is pretty } straightforward to take an image at 0.8 seconds, then blank the beam } very quickly before taking the next image. That way you get pretty close } to the 8 sec total exposure. If there is no beam blanker on the } microscope, in most cases it can be added. } } I am not sure what you mean by "too sensitive". The cameras are usually } constructed so that 1 electron from the beam creates between a few tenth } to a few counts (these are all statistical data, of course). The well } width divided by this sensitivity then determines, how many primary } electrons are needed to fully expose one pixel. For example, if the well } width is 50,000 electrons and the sensitivity is 1 count/electron, one } needs 50,000 primary electrons to fill the well. This translates into } roughly a 0.4% statistical error. } } } From a practical standpoint: You can take images with most cameras when } the exposure meter on the microscope reads a couple of seconds without } overexposing the camera. On the other hand, you can reduce the intensity } of the beam until you see single electron events. } } The one area where CCD cameras may be too sensitive is diffraction. The } normally huge intensity in the transmitted beam often leads to } saturation. In CCDs this can lead to blooming (the intensity spills over } into neighboring pixels). This can be taken care of with special chips } that have anti-blooming features, but this usually has some other } drawbacks. Again, this can also be overcome somewhat with multiple } exposures. Film behaves more civilized here, as it simply stops } responding to the electrons, but this makes film more or less useless } for quantitative measurements of diffraction patterns. I have done } diffraction with CCDs many times and though it does require some } tweaking, one can get very good results from them. } } Michael Bode, Ph.D. } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } =================================== } phone: (888) FIND SIS } (303) 234-9270 } fax: (303) 234-9271 } email: mailto:info-at-soft-imaging.com } web: http://www.soft-imaging.com } =================================== } } } } -----Original Message----- } From: jim [mailto:jim-at-proscitech.com.au] } Sent: Wednesday, May 31, 2000 9:37 PM } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } Subject: RE: Film vs Digital } } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } Yes, it could be done "in theory". Somebody would need to figure out the } } software and perhaps modify the hardware. Then we would find that the } total } exposure of the specimen to the electron beam maybe a muliple of the } film's } exposure. Afterall, an 8 sec film exposure would not amount in digital } to } 10x0.8, but we would require considerable time in between exposures. } Since the } problems in the discussed circumstances are specimen movement and beam } damage, } it seems that taking multiple exposures is a poor option. } } Digital cameras are for some situation too sensitive to electron } exposure. } Cutting back on electrons is no option since its the electrons that form } the } image in the first instance. } Much easier in light microscopy . . . insert a neutral density filter. } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com }
if possible, select plates obtained from serial dilutions which show confluent lysis (i.e., where plaques touch each other). They are a good source for obtaining high-titer bacteriophage lysates (i.g., 10E10 / 10E11 plaque-forming units per ml). Harvest the phage by scraping the top agar and transfer it into a test tube or equivalent. Rinse the plates with some ml of phage diluent (i.g., 5 ml). Shake this supension containing phage, host cells and agar for some while before spinning down the cells and the agar. It is also a good idea to pick up a single plaque. Resuspend it in a small volume of broth, and prepare a fresh lysate in some ml of broth with fresh host cells. For rapid screening, we sometimes pipette a drop of } buffer onto a plaque and float a piece of carbon film into the drop directly from a mica support. After some minutes, we pick up the film with a grid and do the routine negative staining. But you are right, the number of phage particles is low in this case. Best regards Horst Neve
} At 15:14 01.06.00 -0500, you wrote: } Hi, } } We are trying to help a client find and identify a bacteriophage. His } bacteria (Pasturella sp.) are plated out on agar and show well-defined clear } areas with sharp edges where the phage are supposed to be. So far, we have } taken carbon-coated grids and placed them gently onto the surface of the } clear areas, then lifted them off and negative-stained with PTA or uranyl } acetate. We have also run buffer across the clear areas, then pipetted it } onto the grids and stained it. A microbiologist who works with phage in } another lab has taken samples from the clear areas and concentrated them } down and we have stained those also. } } So far we have found exactly two phage-like organisms in a total of about 10 } grids. Not a stellar performance. } } Thanks in advance. } } Randy } } Randy Tindall } EM Specialist } Electron Microscopy Core Facility } W122 Veterinary Medicine } University of Missouri } Columbia, MO 65211
**************************************************************************** **** Dr. Horst Neve Institut fuer Mikrobiologie / Institute for Microbiology Bundesanstalt fuer Milchforschung / Federal Dairy Research Centre Postfach / P.O. Box 6069, D-24121 Kiel Hermann-Weigmann-Str. 1, D-24103 Kiel **************************************************************************** **** Tel. / Phone: +49 (0) 431 609 2343 {} Fax: +49 (0) 431 609 2306 E-mail: neve-at-bafm.de {} Internet: http://www.bafm.de **************************************************************************** ****
if possible, select plates obtained from serial dilutions which show confluent lysis (i.e., where plaques touch each other). They are a good source for obtaining high-titer bacteriophage lysates (i.g., 10E10 / 10E11 plaque-forming units per ml). Harvest the phage by scraping the top agar and transfer it into a test tube or equivalent. Rinse the plates with some ml of phage diluent (i.g., 5 ml). Shake this supension containing phage, host cells and agar for some while before spinning down the cells and the agar. It is also a good idea to pick up a single plaque. Resuspend it in a small volume of broth, and prepare a fresh lysate in some ml of broth with fresh host cells. For rapid screening, we sometimes pipette a drop of } buffer onto a plaque and float a piece of carbon film into the drop directly from a mica support. After some minutes, we pick up the film with a grid and do the routine negative staining. But you are right, the number of phage particles is low in this case. Best regards Horst Neve
} At 15:14 01.06.00 -0500, you wrote: } Hi, } } We are trying to help a client find and identify a bacteriophage. His } bacteria (Pasturella sp.) are plated out on agar and show well-defined clear } areas with sharp edges where the phage are supposed to be. So far, we have } taken carbon-coated grids and placed them gently onto the surface of the } clear areas, then lifted them off and negative-stained with PTA or uranyl } acetate. We have also run buffer across the clear areas, then pipetted it } onto the grids and stained it. A microbiologist who works with phage in } another lab has taken samples from the clear areas and concentrated them } down and we have stained those also. } } So far we have found exactly two phage-like organisms in a total of about 10 } grids. Not a stellar performance. } } Thanks in advance. } } Randy } } Randy Tindall } EM Specialist } Electron Microscopy Core Facility } W122 Veterinary Medicine } University of Missouri } Columbia, MO 65211
**************************************************************************** **** Dr. Horst Neve Institut fuer Mikrobiologie / Institute for Microbiology Bundesanstalt fuer Milchforschung / Federal Dairy Research Centre Postfach / P.O. Box 6069, D-24121 Kiel Hermann-Weigmann-Str. 1, D-24103 Kiel **************************************************************************** **** Tel. / Phone: +49 (0) 431 609 2343 {} Fax: +49 (0) 431 609 2306 E-mail: neve-at-bafm.de {} Internet: http://www.bafm.de **************************************************************************** ****
I'm looking for a (preferably) bench top incubator. Non-water jacketed, not using B.O.D. bottles, unit needs to be using CFC free refrigeration system. Does anyone know of such a unit or where one can be purchased? Need to order one ASAP. We are a small research lab and the one we have is a monster (48"Hx46"Wx28"deep and weighs almost 500lbs.). I'd appreciate any info.
The Microscopy & Microanalysis 2000 Local Arrangements Committee is pleased to announce an additional social event at M&M2000---
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Date sent: Fri, 02 Jun 2000 11:03:03 -0400 To: c.jeffree-at-ed.ac.uk } From: joe fu {jofu-at-nist.gov}
I have a researcher here at USU who would like to have some freeze fracture preps made of lysosome. Is there anyone out there who is currently doing FF preps and would be willing to assist us? I can image the preps here in Logan.
William McManus Supervisor Electron Microscopy Facility Department of Biology Utah State University Logan UT 84322-5305
Having worked with a variety of phages (Candida, Streptococcus, E. coli) I can tell you that finding phages from an agar plaque is very difficult--as you have determined. The best way is to do a liquid culture and then high speed followed by ultracentrifugation to concentrate the particles. FYI, as I recall, on a 200 mesh grid each virus particle is roughly equivalent to 3.4 x 10E6 vp/ml. So, you need a lot of particles to even find one of them using this approach.
JB #################################################################### John J. Bozzola, Ph.D., Director Micro-Imaging and Analysis Center 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 Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
My EM lab is going to be moved and I have just seen the plans. Apparently two large equipment room are going to be built, one right across the hall and the other two doors down. I have been told that the equipment room will house -80 freezers, centrifuges, etc. Does anyone out there have experience with this type of equipment near their TEM? I will not be able to test the room before the move because nothing has been built yet and we all scheduled to move in at the same time. I am hoping to have some influence on the architects now. They have been told (and I shall keep reminding them) that no electrical circuits are to be passed around the EM room.
Thanks.
Ruth
********************************************** Ruth Yamawaki Department of Comparative Medicine Building 330, Quad 7, RAF-1 Stanford CA 94305 (650) 723-3457 **********************************************
Mike Bode would use multiple digital exposures. The exposures could be layered and combined into one superior image. This image would be made up of more pixel and is formed by more electrons and so would be noise-free and hence could be further enlarged then otherwise possible. Perhaps.
No, I did not talk about further enlargements. All I wanted to say is, that a more noise-free image can be achieved by adding multiple images, and that this also to some extent helps with drift of the sample during acquisition.
Jim wrote:
How much time is required between exposures to transfer a minimum 10mb image per exposure?
How did you arrive at 10 MB? A 1280x1024 image with 16 bit pixel information is about 2.5 MB (uncompressed). We acquire about 10 of those per second and transfer them across the PC bus to the display. Putting them on them into Memory might add a few tenth of a second. Writing to HD can be done after all images are acquired.
Jim wrote:
What would be the total time from focusing to the last exposure? What about Z-drift in the interim requiring objective changes
Why would we have to worry about that, if we don't have to worry about that when taking the image on film? In fact, we could take care of this by looking at the image between exposures and correct for z-drift. However, as you said, that would add to the overall time and exposure. I was comparing a normal dark field image taken on film at 8 seconds with acquiring the same image on a "too sensitive" CCD camera by adding up 10 consecutive .8 second images. Why would the sample drift (in x, y or z) substantially more in 8+delta seconds than in 8?
Jim wrote:
what about the total cost of this additional get-up
That of course depends on the microscope and there is no general answer. For example on a LEO 912 I believe the blanker is standard. The additional cost to use an acquisition scheme like this with our software is $0 plus perhaps a bit of time to write a small macro. On other microscopes one might have to add a beam blanker and perhaps a control mechanism for the beam blanker. But I would guess, that this cost is not very high. All modern microscopes are computer controller anyway, so it is most likely just a control command that needs to be sent to the microscope over a serial port if the beam blanker is installed. Piece of cake.
Jim wrote:
The mind boggles at a through focus series.
You're right here. But I don't think we were talking about through-focus series. Incidentally, we do through-focus series on light microscopes and reconstruction routinely. Takes a few images at different focus (or for a light microscope: stage) settings. The rest is done off-line. Takes maybe a couple of minutes for about 20 images of about 1kx1k. I agree that TEM is different here and much more complicated due to the complicated Contrast Transfer Function. However, this could in principle be sorted out.
Jim wrote:
Again, I don't doubt that there is now a large place for digital in TEM, but its no panacea.
I also agree with you on that one. But using the additional computer possibilities of digital imaging might take you further than expected.
Michael
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: jim [mailto:jim-at-proscitech.com.au] Sent: Friday, June 02, 2000 5:15 AM To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' Cc: 'jim-at-proscitech.com.au'
This can be hard to predict. Once we were having trouble with rather huge (100 eV) energy fluctuations in our GIF 200 energy filter. We traced the source to an adjoining room filled with constant-temperature ovens, fans, and other high-current equipment. The unlikely source finally turned out to be a $50 hot plate-stirrer!
Keeping the AC circuitry from running near the lab is a very good start- I know this has devastated other labs. Ask for your own independent electrical ground for the 'scope, and that all electrical circuits to your lab remain independent of other rooms. I doubt that the equipment you describe will be a big problem as long as you don't share AC circuits, ground, or a common wall. Good luck.
"The chief source of problems is solutions." -Eric Sevareid
........................................................................... ...................................................... Jeffrey A. Fortner Argonne National Laboratory CMT/205 9700 S. Cass Avenue Argonne, IL 60439-4837
(630) 252-5594 (voice) (630) 252-4771 (fax)
} ---------- } From: Ruth Yamawaki } Sent: Friday, June 2, 2000 11:48 AM } To: 'Microscopy-at-sparc5.microscopy.com' } Subject: Outside electrical near the EM lab } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } My EM lab is going to be moved and I have just seen the plans. Apparently } two large equipment room are going to be built, one right across the hall } and the other two doors down. I have been told that the equipment room } will } house -80 freezers, centrifuges, etc. Does anyone out there have } experience } with this type of equipment near their TEM? I will not be able to test } the } room before the move because nothing has been built yet and we all } scheduled } to move in at the same time. I am hoping to have some influence on the } architects now. They have been told (and I shall keep reminding them) } that } no electrical circuits are to be passed around the EM room. } } Thanks. } } Ruth } } ********************************************** } Ruth Yamawaki } Department of Comparative Medicine } Building 330, Quad 7, RAF-1 } Stanford CA 94305 } (650) 723-3457 } ********************************************** } }
There is a standard technique for phage isolation and purification in Maniatis. I have found it to work better than any kit and can be modified accordingly.
Horst Neve 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 } -----------------------------------------------------------------------. } } Randy, } } if possible, select plates obtained from serial dilutions which show } confluent lysis (i.e., where plaques touch each other). They are a good } source for obtaining high-titer bacteriophage lysates (i.g., 10E10 / 10E11 } plaque-forming units per ml). Harvest the phage by scraping the top agar } and transfer it into a test tube or equivalent. Rinse the plates with some } ml of phage diluent (i.g., 5 ml). Shake this supension containing phage, } host cells and agar for some while before spinning down the cells and the } agar. It is also a good idea to pick up a single plaque. Resuspend it in a } small volume of broth, and prepare a fresh lysate in some ml of broth with } fresh host cells. For rapid screening, we sometimes pipette a drop of } } buffer onto a plaque and float a piece of carbon film into the drop } directly from a mica support. After some minutes, we pick up the film with } a grid and do the routine negative staining. But you are right, the number } of phage particles is low in this case. } Best regards } Horst Neve } } } At 15:14 01.06.00 -0500, you wrote: } } Hi, } } } } We are trying to help a client find and identify a bacteriophage. His } } bacteria (Pasturella sp.) are plated out on agar and show well-defined clear } } areas with sharp edges where the phage are supposed to be. So far, we have } } taken carbon-coated grids and placed them gently onto the surface of the } } clear areas, then lifted them off and negative-stained with PTA or uranyl } } acetate. We have also run buffer across the clear areas, then pipetted it } } onto the grids and stained it. A microbiologist who works with phage in } } another lab has taken samples from the clear areas and concentrated them } } down and we have stained those also. } } } } So far we have found exactly two phage-like organisms in a total of about 10 } } grids. Not a stellar performance. } } } } Thanks in advance. } } } } Randy } } } } Randy Tindall } } EM Specialist } } Electron Microscopy Core Facility } } W122 Veterinary Medicine } } University of Missouri } } Columbia, MO 65211 } } **************************************************************************** } **** } Dr. Horst Neve } Institut fuer Mikrobiologie / Institute for Microbiology } Bundesanstalt fuer Milchforschung / Federal Dairy Research Centre } Postfach / P.O. Box 6069, D-24121 Kiel } Hermann-Weigmann-Str. 1, D-24103 Kiel } **************************************************************************** } **** } Tel. / Phone: +49 (0) 431 609 2343 {} Fax: +49 (0) 431 609 2306 } E-mail: neve-at-bafm.de {} Internet: http://www.bafm.de } **************************************************************************** } ****
I spent about two years trying to make good pictures of my NanoGold labeled protein-DNA complexes. Doing this job I find two main problems: the sample is unstable under the beam as any biological sample; NanoGold is much more bright than protein core in dark-field. I find that it is impossible to record equally perfect signals from NanoGold and protein core because of short dynamic range for SO-163 film, I believe. I was trying to make two pictures with different exposure, but it is tricky: in dark-field mode the automatic exposure meter usually does not work and we have to set exposure manually, in this case it is difficult to get "right" exposure time in the right moment, you know. Again, because of sample's short life under the beam, it is impossible to make a couple pictures at the different conditions sometime. Keep in mind, please, that to change the film in the microscope it takes about 10 seconds. Your idea about increasing signal-noise ratio by collecting more electrons is bright but not practical. For biological samples (I am talking about non-fixed, non-stained samples of proteins, DNA or RNA-protein complexes etc) the electron damage is a huge problem. People are trying to solve it in different ways. Some using cryo temperature (to stabilize the biological structure). I was using freeze-drying (I find that freeze-dried samples are more stable under the beam). But in any case we have deal with very unstable samples and must to do everything to decrease (not increase as you recommended) electron dose. Drift is a second big problem for such application: to increase signal-noise ratio we have to use very thin support films. Images obtained at such conditions are noisy and in most cases we have to use image analysis tools to extract the data. It means that we have to digitize our images anyway. In such situation digital camera may help. As you, probably, remember I was a person who initiates this discussion. I think this discussion was very useful for many of us who are not friendly with digital camera's techniques. We understand the limitations of the modern digital cameras better now. I would like to say thank you everybody who was involved in this discussion. There is some conclusions I make for myself from discussion:
- Film is still cheap and universal material for recording and storage EM images, sorry CCD. - CCD TEM camera should not substitute film. Film and camera should work all together improving the flexibility of the TEM system. For this reason I will chose side-mount camera if will have money for it. - For cell-biology (thin sections) where the resolution of the sample is about 3 nm CCD camera may do a good job allowing users to make a huge number of pictures (cell-biology guys love it), instantly view and catalogize them. - Sometime the digital camera may help in area of high-resolution (relatively high, guys) EM when image will be digitized anyway. The major limitation here is small area of view (we need a lot of particles for image analysis sometime), but you could make the set of overlapping pictures and digitally combine it. I love, also, Mike Bode idea to make a few very short exposure pictures and combine it digitally later to reduce noise. The relatively big size of CCD's pixels is a real problem too. - CCD camera is expensive "toy". I am not sure that the benefits from using it will compensate astronomical price, actually the third of the electron microscope value ($70000 is it 1/3 of microscope's price on current market?). Currently, I would recognize the CCD TEM camera as funny "attachment" which may be useful if you rich enough to spend money on it (it mean, that you have everything else in your EM lab plus some extra $70000 for the fun playing with digital "toy"). - TEM CCD cameras are under extensive development now. Today's camera will be replaced on the new model (read better, faster, what else?) next year. Next year's camera will be easily forgotten next after the next year and so on... Each new camera will be better that previous one... CCD TEM camera it is not a good investment of money, I think. - We should keep in mind that many companies charged extra 3-4K$ for the installation and training (it is mandatory for Gatan for instance) and you, probably, have to buy service contract on it even if you have service contract on the microscope (JEOL's service contract on microscope do not cover the CCD camera even if you buy it from JEOL).
Best regard, Sergey
} Date: Fri, 02 Jun 2000 21:14:44 +1000 } From: jim {jim-at-proscitech.com.au} } Subject: RE: Film vs Digital } To: 'Mike Bode' {mb-at-Soft-Imaging.com} , } "'Microscopy-at-MSA.Microscopy.Com'" {Microscopy-at-sparc5.microscopy.com} } Cc: "'jim-at-proscitech.com.au'" {jim-at-proscitech.com.au} } Reply-to: "jim-at-proscitech.com.au" {jim-at-proscitech.com.au} } Organization: ProSciTech } X-Mailer: Microsoft Internet E-mail/MAPI - 8.0.0.4211 } } ------------------------------------------------------------------------ } 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
I agree with most of your statements, and I don't think that anybody would argue the point, that a raw CCD chip has a better resolution that film. As you pointed out, film can have a very small grain size ( {1 micron) and CCD chips usually have a few microns pixel size.
But that is not the end of the story. An optical system normally consists of more than a chip or a sheet of film. The question is, can I get the resolution I want or need. And here the situation is not as simple. For example: I used to do high-resolution TEM. What you do there is operate the microscope at optimum condition, then take a picture (on film). You then go to the darkroom and develop prints by blowing up the negative 10, 20 or even more times. When you then look at the images, you can usually see the grains of the film (especially if you then scan those into a copmputer). So, we are working at the resolution limit of the film, and according to your postings, we should not be able to see anything on a CCD. But that's not true. By using some geometrical properties (the camera sits further down in the column and sees an already enlarged image) and a tapered fiber-optic, you can acquire just as good and better images of the same structure. Both images are limited by the point resolution of the TEM and not by the Film or CCD resolution.
What I am trying to say, and what I have said before is, that film is definitely better when it comes to maximizing the product of resolution AND field of view. However, if we can trade one for the other, I believe in most cases you get better results from a CCD.
Having said that and looking at a print of Ansel Adams, I am glad there is film, though!!
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: Chris Jeffree [mailto:cjeffree-at-srv0.bio.ed.ac.uk] Sent: Friday, June 02, 2000 10:16 AM To: joe fu Cc: microscopy-at-sparc5.microscopy.com
Date sent: Fri, 02 Jun 2000 11:03:03 -0400 To: c.jeffree-at-ed.ac.uk } From: joe fu {jofu-at-nist.gov}
Conventional wisdom said that when it came to using TEM for biological specimens (or beam sensitive specimens) it is always best to use a lower accelerating voltage. Many lower voltage microscopes (V { 120 kV) were sold on this assumption. However is this assumption true? At the present there are many research establishments, buying TEMs for biological use, who are using 200 to 300 kV beams. So obviously there has been a shift in the conventional way of thinking. Being materials based I am not sure what the status quo is in biological TEM. I know that the ratio of inelastic to elastic scattering cross sections is greater than one for the elements Z {12, but how does this change as the beam energy increases? What are your experiences? This is really just academic curiosity by the way, but I am sure many of you would appreciate the question. ******************************************************** Dr Jonathan Barnard
Analytical Materials Physics The Angstrom Laboratory, Uppsala University P O Box 534, SE-751 21 Uppsala, Sweden Phone: +46-(0)18-4716838 Fax: +46-(0)18-500131 Phone: Microscope room +46 18 471 6365 http://www.angstrom.uu.se/analytical/home.html ********************************************************
} } - CCD camera is expensive "toy". I am not sure that the } } benefits from using it will compensate astronomical price, actually } } the third of the electron microscope value ($70000 is it 1/3 of } } microscope's price on current market?). Currently, I would } } recognize the CCD TEM camera as funny "attachment" which may be } } useful if you rich enough to spend money on it (it mean, that you } } have everything else in your EM lab plus some extra $70000 for the } } fun playing with digital "toy"). - TEM CCD cameras are under extensive development now. Today's camera will be replaced on the new model (read better, faster, what else?) next year. Next year's camera will be easily forgotten next after the next year and so on... Each new camera will be better that previous one... CCD TEM camera it is not a good investment of money, I think. { {
Sergey:
I appreciate your post, but...
1. Remember that "time is money"...there is no question that there is a value in the nearly instantaneous result that derives from the use of digital imaging systems, particular on TEMs. Also, the ability to rapidly process and analyze your images to determine if they are "keepers" is priceless, IMO.
2. In our large, national multi-user facility we have been digital-only since about 1993 or so. We have no darkrooms for plate loading or enlarging. The only "chemicals" we handle in the entire photographic process are toner cartridges for our laser printers, or equivalent media for dye sub printers etc. In most instances, we process our images electronically all the way through to the final presentation. This includes preparation of PowerPoint slides for digital projection for talks, to sending full papers out for publication on disks or via e-mail attachments. No user has *ever* complained about the non-availability of film, or about the "lack of resolution of CCD images" compared to film. On the contrary, we have users that travel to our laboratory specifically to do work on our instruments because of the availability of digital imaging, when the same instruments in their own laboratories are not equipped with digital cameras.
3. Digital camera systems also provide the capability to conduct research with outside colleagues live-time via Telepresence Microscopy methods. This is a rapidly developing capability in our field that just cannot be done (on a TEM at least) if your microsocope only uses film for imaging.
4. The 1k x 1k Gatan CCD camera on our Hitachi HF-2000 was purchased for about $100K in 1993, and was upgraded about 5 years ago to a multi-scan capability. It is still functioning perfectly today, as is a similar camera on our JEOL 4000EX TEM. The very newest CCD cameras might offer faster read-out times, but there has been no order-of-magnitude improvement in capabilities to make our older camera so obsolete that we desire to purchase a new one. The point is that, unlike desk-top computers, the *expensive* digital camera you purchase today will definitely *not* be obsolete next year...
5. A CCD TEM camera is probably the *best* investment anyone could make to advance the research throughput in their microscopy facility.
All just MHO...
Larry
Dr. Lawrence F. Allard Senior Research Staff Member High Temperature Materials Laboratory Oak Ridge National Laboratory 1 Bethel Valley Road Bldg. 4515, MS 6064 PO Box 2008 Oak Ridge, TN 37831-6064
} From: "Mike Bode" {mb-at-Soft-Imaging.com} To: "'Microscopy-at-MSA.Microscopy.Com'" {Microscopy-at-sparc5.microscopy.com} } Chris, } } I agree with most of your statements, and I don't think that anybody } would argue the point, that a raw CCD chip has a better resolution that } film. As you pointed out, film can have a very small grain size ( {1 } micron) and CCD chips usually have a few microns pixel size. } } But that is not the end of the story. An optical system normally } consists of more than a chip or a sheet of film. The question is, can I } get the resolution I want or need. And here the situation is not as } simple. For example: I used to do high-resolution TEM. What you do there } is operate the microscope at optimum condition, then take a picture (on } film). You then go to the darkroom and develop prints by blowing up the } negative 10, 20 or even more times. When you then look at the images, } you can usually see the grains of the film (especially if you then scan } those into a computer). So, we are working at the resolution limit of } the film, and according to your postings, we should not be able to see } anything on a CCD. But that's not true. By using some geometrical } properties (the camera sits further down in the column and sees an } already enlarged image) and a tapered fiber-optic, you can acquire just } as good and better images of the same structure. Both images are limited } by the point resolution of the TEM and not by the Film or CCD } resolution.c
For a scanning device be it light, electrons, x-rays or gamma rays is a CCD array the best way to capture the image. My experience has been with gamma rays and to some extent x rays. We have not found a good way to focus gamma rays so we use a crystal that emitted light and a photomultiplier tube and we got good images. The resolution depended on the aperture of the gamma ray source and was pretty large.
If you are scanning a sample wiht an electron beam the same principle should work. You would not need long dwell times but build the images out of multiple scans.
If you are scanning the sample the array of pixels seems redundant to me. Also a photomultiplier tube and crystal are a great deal more sensitive than CCD arrays and have a good deal more dynamic range.
The time to make a film image is allows going to be less than making a digital image. The ability to immediately see the digital image is a very handy thing. There are ways to develop B&W film that you can see your image in 5 minutes. For 4 X 5 images I am using BZT tubes that are tubs with a inter circumference of a little over 4 inches. You put the file and developer in the tube and put the tub in a pan of water and spin the tube in the water. The stop bath and fixing can be carried out in room light and it results in the most even development I have ever seen.
For archival storage I don't think 35 mm film can be beat for cost and resolution. Unfortunately you can't see the results until later occasional making it necessary to reshoot the session if you rely on film alone.
If you really need long term archival storage you might consider using a slide printer for a computer to print the digital images. We know the life of silver film is longer then 100 years. For the LM folks that are using color images you could use PhotoShop to do a color separation and print out three negatives on black and white for storage.
Hard drives And CD ROMS have come a long way but I loose about 20% of my hard drive storage a year and loose an occasional CDROM to scratches. Film would survive the scratches with a minimal loss of information insted loosing the whole CDROM.
Gordon
Gordon Couger gcouger-at-couger.com
Stillwater, OK www.couger.com/gcouger 405 624-2855 GMT -6:00
Please be aware that this is a commercial post that may be of interest to some of the list members.
Digital light microscope cameras are now available that use a highly touted CCD detector: Color or monochrome; fast interline transfer, high sensitivity, low noise; 6.7 micron square pixel; 2/3 inch 1.3 Million chip; PCI bus frame transfer. This detector is ideal to use in light microscopy applications. Now precision technology and fast high capacity cheap computers allow this detector to be "Micro Stepped" providing digital images of up to 12 million pixels to be captured very quickly and providing file sizes of up to 35 MB in 24 bit color images. "Inter Pixel Stepping" technology will allow a considerably less expensive approach to digital imaging that can now approach film resolving capability in a light microscope.
Hats of to Dr. Jeffree's explanation on optical resolution as it relates to light microscopy applications. With high pixel density capable digital cameras, now lower magnification, low NA (lower resolving power) objectives can be used to acquire matching digital resolution images of wide fields of view.
Information on Nikon's Instrument Division "Digital Eclipse" family of digital cameras for microscopy including the new DXM1200 digital color camera will soon be available on our web site www.nikonusa.com or you can go to our Dealer Locator at http://www.nikonusa.com/corpinfo/dealers/dealerSearch.cfm and ask your local Nikon authorized microscope dealer for further information.
Best Regards,
Stan Schwartz Manager, BioSciences Dept. Nikon Inc Instrument Division 1300 Walt Whitman Rd. Melville, NY 11747 631-547-8500 631-547-4033 Fax Schwartz-at-nikonincmail.com www.nikonusa.com
Earlier post ============================================================= Geoff Analysing this a little further: A 25 Mbyte image = 8.3 M pixels to cover 24x36mm or 864mm^2 this represents 98.2 pixels per mm, or 49 line pairs per mm. Pixel size 10.2 um. Resolution about 50 line pairs per mm at best.
A 25M pixels image used to capture a 24x36mm Kodachrome slide represents 28,935 pixels per mm^2 or 170 pixels per mm. This is equivalent to a maximum resolution of 85 line pairs per mm, which may be on the conservative side for Kodachrome. pixel size = 5.88um. The image will be approx. 6120x4082 pixels, generating a file size of approx 75Mb for a 24-bit (8+8+8bit) RGB image.
To record 120 line pairs per mm, which many top 35mm camera lenses can achieve, a minimum of 240 pixels per mm are required, each 4.2 um wide. This equates to 8640x5760 pixels for a 24x36mm frame = 50M-pixels or 50Mb in 8-bit greyscale, or 150Mb in 24-bit RGB.
At 320 line pairs per mm (Technical Pan) the minimum required 640 pixels per mm is a pixel size of 1.56 um
Presumably for a light image the diffraction limited resolution is approx 1/2 lambda which at 540nm is 0.27um. So looking to the future of ultimate-performance CCDs, direct recording of a diffraction limited light image projected onto the sensor requires at the very least 3703 pixels per image mm or 13,717,421 pixels per mm^2 (greyscale 8-bit)
However, if we are doing light microscopy with an NA 1.4 x100 objective, how much resolving power do we need on CCD or film?
Data is at 0.27um resolution (lambda = 540nm). Let's round this to 0.3um. Magnification at 24x36mm film image is x100, so pixels must be an absolute maximum of 30um wide to record the significant data = 33.3 pixels per mm, equivalent to 800x1200 pixels to record the whole 35mm negative area. However, most CCDs are much smaller than 35 mm frames, typically 1/3 inch. So an 800x1200 pixel CCD at 8x12 mm, 1/3 of the linear dimensions of a 35mm frame would use 9 pixels to record the smallest image details. This is about right from the point of view of resolution, but to record the whole 35mm frame we need about 2400x3600 pixels on our CCD.
Note also that Technical Pan has (depending on the criterion used to assess its performance) up to 10 times the resolving power required to record all there is to see in a diffraction-limited LM image made with a 100x NA 1.4 lens. So you can comfortably afford to use a 60x NA 1.4 lens, thereby getting the same resolution with a bigger field of view.
Many years ago, I took a photograph of a street scene using a Canon 35mm SLR loaded with Kodak Recordak (I think this was a single layer microfilm emulsion). Examined in a light microscope, the image clearly, legibly recorded the brand-name of a child's push chair. I tried to print this brand name using a DeVere point- source enlarger with an image size of 20x30 inches produced with a Schneider Componon lens, but was completely unable to produce a legible image. The point I am making here is that the combination of some high performance films, with high quality lenses of the standard produced by the leading camera manufacturers can record more detail on the film than you can easily get back out by conventional printing. I suspect the same is true of EM exposures.
So there is no contest - film beats CCDs for resolution hands down. And you can process the image on the cheap. No money goes to Intel or Microscoft, Adobe or Epson. But resolution is not primarily what we buy CCDs for. We buy them primarily for instant image capture in a format suitable for digital storage, digital transmission quantitative data recording and image processing.
Chris
===================================================================== DR CHRIS JEFFREE BIOSEM - BIOLOGICAL SCIENCES EM FACILITY UNIVERSITY OF EDINBURGH Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 131 650 5345 FAX. #44 131 650 6563 Mobile 0410 585 401 email c.jeffree-at-ed.ac.uk SEM / TEM bookings sem-at-ed.ac.uk =====================================================================
I work with EPON sections of biological specimens. It seems clear to me that 60KV introduces more dammage than 80Kv. The sections are more unstable and tend to break more easily at the lower voltage.
Dr. A.P. Alves de Matos Pathology Department Curry Cabral Hospital Lisbon
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Conventional wisdom said that when it came to using TEM for biological specimens (or beam sensitive specimens) it is always best to use a lower accelerating voltage. Many lower voltage microscopes (V { 120 kV) were sold on this assumption. However is this assumption true? At the present there are many research establishments, buying TEMs for biological use, who are using 200 to 300 kV beams. So obviously there has been a shift in the conventional way of thinking. Being materials based I am not sure what the status quo is in biological TEM. I know that the ratio of inelastic to elastic scattering cross sections is greater than one for the elements Z {12, but how does this change as the beam energy increases? What are your experiences? This is really just academic curiosity by the way, but I am sure many of you would appreciate the question. ******************************************************** Dr Jonathan Barnard
Analytical Materials Physics The Angstrom Laboratory, Uppsala University P O Box 534, SE-751 21 Uppsala, Sweden Phone: +46-(0)18-4716838 Fax: +46-(0)18-500131 Phone: Microscope room +46 18 471 6365 http://www.angstrom.uu.se/analytical/home.html ********************************************************
At 02:21 PM 6/3/00, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I hope not.
} Digital light microscope cameras are now available that use a highly touted } CCD detector: Color or monochrome; fast interline transfer, high sensitivity, } low noise; 6.7 micron square pixel; 2/3 inch 1.3 Million chip; PCI bus frame } transfer. This detector is ideal to use in light microscopy applications. } Now precision technology and fast high capacity cheap computers allow this } detector to be "Micro Stepped" providing digital images of up to 12 million } pixels to be captured very quickly and providing file sizes of up to 35 MB in } 24 bit color images. "Inter Pixel Stepping" technology will allow a } considerably less expensive approach to digital imaging that can now approach } film resolving capability in a light microscope. } } Hats of to Dr. Jeffree's explanation on optical resolution as it relates to } light microscopy applications. With high pixel density capable digital } cameras, now lower magnification, low NA (lower resolving power) objectives } can be used to acquire matching digital resolution images of wide fields of } view. } } Information on Nikon's Instrument Division "Digital Eclipse" family of } digital cameras for microscopy including the new DXM1200 digital color camera } will soon be available on our web site www.nikonusa.com or you can go to } our Dealer Locator at } http://www.nikonusa.com/corpinfo/dealers/dealerSearch.cfm and ask your } local Nikon authorized microscope dealer for further information. } } Best Regards, } } Stan Schwartz } Manager, BioSciences Dept. } Nikon Inc Instrument Division } 1300 Walt Whitman Rd. } Melville, NY 11747 } 631-547-8500 } 631-547-4033 Fax } Schwartz-at-nikonincmail.com } www.nikonusa.com
As a long term past user of Nikon equipment, it is a tale of sorrow. Most recently, the digital E1 and E2 are dismal failures. Consumers are sending back 990's in droves (see rec.photo.marketplace.digital). I really think that Nikon blew it in regards to digicams. How Nikon can claim to get a 1.3M pixel imager to produce realistic 12M pixel images is rather absurd....if not offensive. The most recent D1 is a joke. Albeit, an expensive one.
Nikon blew it in the scanner arena and made huge blunders in high end pro digicams (E1 & E2). Total junk from my personal experiences. I am not at all prone to spend a dime on any new Nikon digital things. In fact, I have dumped my Nikon so-called pro lenses and bodies for Contax. But this is another story.
My advice is to be very wary....be very wary of Nikon. I do not use Nikon cameras anymore and I do not use Nikon microscopes anymore. But it is your money and your decision. As they say, "Caveat emptor."
gg
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Modern surfers use PC boards. You can too at http://photoweb.net ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Rubbish! With all due respect this is a complete distortion and mis- use of the point I made. To realise this objective with low NA lenses you would have to find some way to defeat the laws of physics. No CCD imager, however clever will make it possible to correct the shortcomings of cheap, low-performance optics. You clearly misunderstood the point, which is that microscope lens resoltuion is not fundamentally dependent on the magnification factor but on the numerical aperture, which also determines resolution. A x100 NA 1.4 lens resolves no more detail than a x60 NA 1.4, but simply magnifies the image further. This is "empty magnification", which is only useful if your image sensor (film or CCD) has limited resolving power. With a very high resolution film like Technical Pan, you can take advantage of this fact to capture a wide-field diffraction-limited image with a x60 1.4 lens. That is not an option with a low NA lens irrespective of the properties of the sensor, CCD or otherwise.
} Hats of to Dr. Jeffree's explanation on optical resolution as it relates to } light microscopy applications. With high pixel density capable digital } cameras, now lower magnification, low NA (lower resolving power) objectives } can be used to acquire matching digital resolution images of wide fields of } view. } } Information on Nikon's Instrument Division "Digital Eclipse" family of } digital cameras for microscopy including the new DXM1200 digital color camera } will soon be available on our web site www.nikonusa.com or you can go to } our Dealer Locator at } http://www.nikonusa.com/corpinfo/dealers/dealerSearch.cfm and ask your } local Nikon authorized microscope dealer for further information. } } Best Regards, } } Stan Schwartz } Manager, BioSciences Dept. } Nikon Inc Instrument Division } 1300 Walt Whitman Rd. } Melville, NY 11747 } 631-547-8500 } 631-547-4033 Fax } Schwartz-at-nikonincmail.com } www.nikonusa.com } } } Earlier post } ============================================================= } Geoff } Analysing this a little further: } A 25 Mbyte image = 8.3 M pixels to cover 24x36mm or 864mm^2 } this represents 98.2 pixels per mm, or 49 line pairs per mm. Pixel } size 10.2 um. Resolution about 50 line pairs per mm at best. } } A 25M pixels image used to capture a 24x36mm Kodachrome } slide represents 28,935 pixels per mm^2 or 170 pixels per mm. } This is equivalent to a maximum resolution of 85 line pairs per mm, } which may be on the conservative side for Kodachrome. pixel size } = 5.88um. The image will be approx. 6120x4082 pixels, generating } a file size of approx 75Mb for a 24-bit (8+8+8bit) RGB image. } } To record 120 line pairs per mm, which many top 35mm camera } lenses can achieve, a minimum of 240 pixels per mm are required, } each 4.2 um wide. This equates to 8640x5760 pixels for a } 24x36mm frame = 50M-pixels or 50Mb in 8-bit greyscale, or 150Mb } in 24-bit RGB. } } At 320 line pairs per mm (Technical Pan) the minimum required } 640 pixels per mm is a pixel size of 1.56 um } } Presumably for a light image the diffraction limited resolution is } approx 1/2 lambda which at 540nm is 0.27um. } So looking to the future of ultimate-performance CCDs, direct } recording of a diffraction limited light image projected onto the } sensor requires at the very least 3703 pixels per image mm or } 13,717,421 pixels per mm^2 (greyscale 8-bit) } } However, if we are doing light microscopy with an NA 1.4 x100 } objective, how much resolving power do we need on CCD or film? } } Data is at 0.27um resolution (lambda = 540nm). Let's round this to } 0.3um. Magnification at 24x36mm film image is x100, so pixels } must be an absolute maximum of 30um wide to record the } significant data = 33.3 pixels per mm, equivalent to 800x1200 } pixels to record the whole 35mm negative area. However, most } CCDs are much smaller than 35 mm frames, typically 1/3 inch. So } an 800x1200 pixel CCD at 8x12 mm, 1/3 of the linear dimensions } of a 35mm frame would use 9 pixels to record the smallest image } details. This is about right from the point of view of resolution, but } to record the whole 35mm frame we need about 2400x3600 pixels } on our CCD. } } Note also that Technical Pan has (depending on the criterion used } to assess its performance) up to 10 times the resolving power } required to record all there is to see in a diffraction-limited LM } image made with a 100x NA 1.4 lens. So you can comfortably } afford to use a 60x NA 1.4 lens, thereby getting the same } resolution with a bigger field of view. } } Many years ago, I took a photograph of a street scene using a } Canon 35mm SLR loaded with Kodak Recordak (I think this was a } single layer microfilm emulsion). Examined in a light microscope, } the image clearly, legibly recorded the brand-name of a child's } push chair. I tried to print this brand name using a DeVere point- } source enlarger with an image size of 20x30 inches produced with } a Schneider Componon lens, but was completely unable to } produce a legible image. The point I am making here is that the } combination of some high performance films, with high quality } lenses of the standard produced by the leading camera } manufacturers can record more detail on the film than you can } easily get back out by conventional printing. I suspect the same is } true of EM exposures. } } So there is no contest - film beats CCDs for resolution hands } down. And you can process the image on the cheap. No money } goes to Intel or Microscoft, Adobe or Epson. But resolution is not } primarily what we buy CCDs for. We buy them primarily for instant } image capture in a format suitable for digital storage, digital } transmission quantitative data recording and image processing. } } Chris } } ===================================================================== } DR CHRIS JEFFREE } BIOSEM - BIOLOGICAL SCIENCES EM FACILITY } UNIVERSITY OF EDINBURGH } Daniel Rutherford Building } King's Buildings, Mayfield Road } EDINBURGH, EH9 3JH, Scotland, UK } Tel. #44 131 650 5345 } FAX. #44 131 650 6563 } Mobile 0410 585 401 } email c.jeffree-at-ed.ac.uk } SEM / TEM bookings sem-at-ed.ac.uk } ===================================================================== }
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Dr Chris Jeffree University of Edinburgh Biological Sciences EM Facility Daniel Rutherford Building King's Buildings EDINBURGH EH9 3JH Tel: +44 (0) 131 650 5345 FAX: +44 (0) 131 650 6563
Lets place the exhausted and exhausting Digital topic aside. Sergey, you do have a challenging project and one that is worth discussing. Just maybe somebody has an idea that will help you. I think that we discuss in this forum pixels too much and microscopy too little.
You probably have tried most of my following suggestions, but just incase, here are a couple of my thoughts:
Dark field contrast is enhanced most when the density between specimen and background is greatest. So its most effective with no specimen support. You could try your luck with the superfine mesh grids now available; these thin bar grids can be purchased down to 2000 mesh and these grids have a 7.5um hole size. If that is not a fine enough support, then try holey films with a net-like structure.
Carbon coating will stabilize samples dramatically. A little loss of contrast is inevitable, but a least carbon on the grid or holey plastic film does not matter.
Don't see why you want to render the gold nano particles with detail (I assume some greys). In TEM I would expect gold above about 5nm to be black. In STEM or FESEM you could get greys easily, but you may not get the resolution required.
Increasing if available to 200 or more the kV, will give more brightness, less contrast, better resolution (specimen dependent). Most importantly, specimen damage frequently is less at higher kV, that depends on the specimen again. I think that damage is less in specimens with greater electron transparency, but one of our "physical" gurus may explain the whys and wherefores.
Without switching film-types, you can develop the TEM films in things like Microdol-X, Microphen or Rodinal (hope those developers still exist). You could also use D-19 more dilute than normal. For most EM purposes all of these would give too softer negatives, but they just may suit you. Over developing in light photography yields more contrast - not so in TEM, where more electrons are the main contrast mechanism. Hence slower film and denser negatives are preferred, especially by most biologists.
I think that we started out with too much contrast, but a grainy image because of insufficient electron exposure. Hmmm, I think that we should take a holiday. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Saturday, June 03, 2000 7:44 AM, Sergey Ryazantsev [SMTP:sryazant-at-ucla.edu] wrote: } } } Dear Jim } } I spent about two years trying to make good pictures of my NanoGold labeled } protein-DNA complexes. Doing this job I find two main problems: the } sample is unstable under the beam as any biological sample; NanoGold is } much more bright than protein core in dark-field. I find that it is } impossible to record equally perfect signals from NanoGold and protein core } because of short dynamic range for SO-163 film, I believe. I was trying to } make two pictures with different exposure, but it is tricky: in dark-field } mode the automatic exposure meter usually does not work and we have to set } exposure manually, in this case it is difficult to get "right" exposure } time in the right moment, you know. Again, because of sample's short life } under the beam, it is impossible to make a couple pictures at the different } conditions sometime. Keep in mind, please, that to change the film in the } microscope it takes about 10 seconds. Your idea about increasing } signal-noise ratio by collecting more electrons is bright but not } practical. For biological samples (I am talking about non-fixed, } non-stained samples of proteins, DNA or RNA-protein complexes etc) the } electron damage is a huge problem. People are trying to solve it in } different ways. Some using cryo temperature (to stabilize the biological } structure). I was using freeze-drying (I find that freeze-dried samples } are more stable under the beam). But in any case we have deal with very } unstable samples and must to do everything to decrease (not increase as you } recommended) electron dose. Drift is a second big problem for such } application: to increase signal-noise ratio we have to use very thin } support films. Images obtained at such conditions are noisy and in most } cases we have to use image analysis tools to extract the data. It means } that we have to digitize our images anyway. In such situation digital } camera may help. As you, probably, remember I was a person who initiates } this discussion. I think this discussion was very useful for many of us } who are not friendly with digital camera's techniques. We understand the } limitations of the modern digital cameras better now. I would like to say } thank you everybody who was involved in this discussion. There is some } conclusions I make for myself from discussion: } } } - Film is still cheap and universal material for recording and } storage EM images, sorry CCD. } - CCD TEM camera should not substitute film. Film and camera should } work all together improving the flexibility of the TEM system. For this } reason I will chose side-mount camera if will have money for it. } - For cell-biology (thin sections) where the resolution of the sample } is about 3 nm CCD camera may do a good job allowing users to make a huge } number of pictures (cell-biology guys love it), instantly view and } catalogize them. } - Sometime the digital camera may help in area of high-resolution } (relatively high, guys) EM when image will be digitized anyway. The major } limitation here is small area of view (we need a lot of particles for image } analysis sometime), but you could make the set of overlapping pictures and } digitally combine it. I love, also, Mike Bode idea to make a few very } short exposure pictures and combine it digitally later to reduce noise. The } relatively big size of CCD's pixels is a real problem too. } - CCD camera is expensive "toy". I am not sure that the benefits from } using it will compensate astronomical price, actually the third of the } electron microscope value ($70000 is it 1/3 of microscope's price on } current market?). Currently, I would recognize the CCD TEM camera as funny } "attachment" which may be useful if you rich enough to spend money on it } (it mean, that you have everything else in your EM lab plus some extra } $70000 for the fun playing with digital "toy"). } - TEM CCD cameras are under extensive development now. Today's } camera will be replaced on the new model (read better, faster, what else?) } next year. Next year's camera will be easily forgotten next after the next } year and so on... Each new camera will be better that previous one... CCD } TEM camera it is not a good investment of money, I think. } - We should keep in mind that many companies charged extra 3-4K$ for } the installation and training (it is mandatory for Gatan for instance) and } you, probably, have to buy service contract on it even if you have service } contract on the microscope (JEOL's service contract on microscope do not } cover the CCD camera even if you buy it from JEOL). } } Best regard, Sergey } } } } } } Date: Fri, 02 Jun 2000 21:14:44 +1000 } } From: jim {jim-at-proscitech.com.au} } } Subject: RE: Film vs Digital } } To: 'Mike Bode' {mb-at-Soft-Imaging.com} , } } "'Microscopy-at-MSA.Microscopy.Com'" {Microscopy-at-sparc5.microscopy.com} } } Cc: "'jim-at-proscitech.com.au'" {jim-at-proscitech.com.au} } } Reply-to: "jim-at-proscitech.com.au" {jim-at-proscitech.com.au} } } Organization: ProSciTech } } X-Mailer: Microsoft Internet E-mail/MAPI - 8.0.0.4211 } } } } ------------------------------------------------------------------------ } } 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 } } -----------------------------------------------------------------------. } } } } } } The world is full of possible solutions, but are they practical.? } } To produce high-resolution, dark-field or any others TEM images that require } } more electrons to form a clear image, Mike Bode would use multiple digital } } exposures. The exposures could be layered and combined into one superior } } image. } } This image would be made up of more pixel and is formed by more electrons } } and } } so would be noise-free and hence could be further enlarged then otherwise } } possible. Perhaps. } } Beam blanking would largely save the specimen from beam damage and drift } } could } } be compensated for by matching up the digitals. Great. } } How much time is required between exposures to transfer a minimum 10mb image } } per exposure? What would be the total time from focusing to the last } } exposure? } } What about Z-drift in the interim requiring objective changes and what about } } the total cost of this additional get-up. The mind boggles at a through } } focus } } series. } } When pushing the limits a piece of film seems more effective, cheaper and fa } } ster. } } Again, I don't doubt that there is now a large place for digital in TEM, but } } its no panacea. } } Cheers } } Jim Darley } } ProSciTech Microscopy PLUS } } PO Box 111, Thuringowa QLD 4817 Australia } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } www.proscitech.com } } } } On Friday, June 02, 2000 1:24 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] wrote: } } } } } } } } No, it's not really a problem. It's been done with low density } } } microscopy all the time. Granted, there are some technical aspects to be } } } overcome, but (and I can only speak for ourselves) we have done that on } } } a number of microscopes. You are of course correct, that 10 images at } } } 0.8 seconds take longer than 8 seconds as the image has to be } } } transferred, etc. BUT: that's what beam blankers are for. It is pretty } } } straightforward to take an image at 0.8 seconds, then blank the beam } } } very quickly before taking the next image. That way you get pretty close } } } to the 8 sec total exposure. If there is no beam blanker on the } } } microscope, in most cases it can be added. } } } } } } I am not sure what you mean by "too sensitive". The cameras are usually } } } constructed so that 1 electron from the beam creates between a few tenth } } } to a few counts (these are all statistical data, of course). The well } } } width divided by this sensitivity then determines, how many primary } } } electrons are needed to fully expose one pixel. For example, if the well } } } width is 50,000 electrons and the sensitivity is 1 count/electron, one } } } needs 50,000 primary electrons to fill the well. This translates into } } } roughly a 0.4% statistical error. } } } } } } } From a practical standpoint: You can take images with most cameras when } } } the exposure meter on the microscope reads a couple of seconds without } } } overexposing the camera. On the other hand, you can reduce the intensity } } } of the beam until you see single electron events. } } } } } } The one area where CCD cameras may be too sensitive is diffraction. The } } } normally huge intensity in the transmitted beam often leads to } } } saturation. In CCDs this can lead to blooming (the intensity spills over } } } into neighboring pixels). This can be taken care of with special chips } } } that have anti-blooming features, but this usually has some other } } } drawbacks. Again, this can also be overcome somewhat with multiple } } } exposures. Film behaves more civilized here, as it simply stops } } } responding to the electrons, but this makes film more or less useless } } } for quantitative measurements of diffraction patterns. I have done } } } diffraction with CCDs many times and though it does require some } } } tweaking, one can get very good results from them. } } } } } } Michael Bode, Ph.D. } } } Soft Imaging System Corp. } } } 1675 Carr St., #105N } } } Lakewood, CO 80215 } } } =================================== } } } phone: (888) FIND SIS } } } (303) 234-9270 } } } fax: (303) 234-9271 } } } email: mailto:info-at-soft-imaging.com } } } web: http://www.soft-imaging.com } } } =================================== } } } } } } } } } } } } -----Original Message----- } } } From: jim [mailto:jim-at-proscitech.com.au] } } } Sent: Wednesday, May 31, 2000 9:37 PM } } } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } } } Subject: RE: Film vs Digital } } } } } } } } } ------------------------------------------------------------------------ } } } 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 } } } -----------------------------------------------------------------------. } } } } } } } } } Yes, it could be done "in theory". Somebody would need to figure out the } } } } } } software and perhaps modify the hardware. Then we would find that the } } } total } } } exposure of the specimen to the electron beam maybe a muliple of the } } } film's } } } exposure. Afterall, an 8 sec film exposure would not amount in digital } } } to } } } 10x0.8, but we would require considerable time in between exposures. } } } Since the } } } problems in the discussed circumstances are specimen movement and beam } } } damage, } } } it seems that taking multiple exposures is a poor option. } } } } } } Digital cameras are for some situation too sensitive to electron } } } exposure. } } } Cutting back on electrons is no option since its the electrons that form } } } the } } } image in the first instance. } } } Much easier in light microscopy . . . insert a neutral density filter. } } } Cheers } } } Jim Darley } } } ProSciTech Microscopy PLUS } } } PO Box 111, Thuringowa QLD 4817 Australia } } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } } www.proscitech.com } } } } } _____________________________________ } } Sergey Ryazantsev Ph. D. } Electron Microscopy } UCLA School of Medicine } Department of Biological Chemistry } Box 951737 } Los Angeles, CA 90095-1737 } } Phone: (310) 825-1144 } Pager: (310) 845-0248 } FAX (departmental): (310) 206-5272 } mailto:sryazant-at-ucla.edu } http://www.bol.ucla.edu/~sryazant } }
I have two replies and Mike my well have four replies to these. 1. There has been a parallel discussion concerning resolution of film versus digital images. Clearly in raw power digital cannot compete since film has multi gigabyte capacity. I added to that thread that what matters is: does digital have enough power and that frequently it would. Mike reinforced and strengthened that argument, finishing with the note that he is glad for film when looking at prints by Ansel Adams. (Ansel Adams until about 30 years ago carted for decades large format cameras through US National Parks, especially Yosemite, producing fantastic landscape photographs and books) Adams' limited edition prints were contacts of 5x7 and 8x10" inch sheet film, probably rated at 400 ISO. The line resolution of such prints much exceeds our eyes' resolution, but still results in superior gradation and detail. TEM film, even when much enlarged has such details too. Why Mike, should we accept 2.5 Mb? That is a splendid file size for SEM, and because of the limited enlargability of light microscopy (concrete ceiling due to wavelengths) it's reasonable, but minimal for light microscopy. TEM can do and deserves better.
Incidentally, from the outset I cited increased "enlargability" to obtain high resolution TEM, as one of films major advantages, since greater depths of field at moderate powers makes high powers through photo enlarging a desirable technique. The small additional magnification yielded by placing a digital camera lower in the column does not compensate. So greater "enlargability" of digitals would be desirable, but is limited by pixel size.
2 The thread was initiated by Sergey. He had problems visualising certain specimens in dark field. The use of his "beaut" digital TEM camera made things worse. I pointed out that the shorter exposure reduced the number of electrons forming the image, hence more noise. I believe that a good part of Mike's case will be settled in his favour when we hear "Eureka" from Sergey's lab. I don't doubt that much more can be done with digital now and that further improvements are on the way. Mike's "solution" may well be possible, but I don't believe its a snap; in any case: Show Sergey! Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Saturday, June 03, 2000 2:58 AM, Mike Bode [SMTP:mb-at-Soft-Imaging.com] wrote: } } Well, let's see: } } Jim wrote: } } Mike Bode would use multiple digital } exposures. The exposures could be layered and combined into one superior } image. } This image would be made up of more pixel and is formed by more } electrons and } so would be noise-free and hence could be further enlarged then } otherwise } possible. Perhaps. } } No, I did not talk about further enlargements. All I wanted to say is, } that a more noise-free image can be achieved by adding multiple images, } and that this also to some extent helps with drift of the sample during } acquisition. } } Jim wrote: } } How much time is required between exposures to transfer a minimum 10mb } image } per exposure? } } How did you arrive at 10 MB? A 1280x1024 image with 16 bit pixel } information is about 2.5 MB (uncompressed). We acquire about 10 of those } per second and transfer them across the PC bus to the display. Putting } them on them into Memory might add a few tenth of a second. Writing to } HD can be done after all images are acquired. } } Jim wrote: } } What would be the total time from focusing to the last exposure? } What about Z-drift in the interim requiring objective changes } } Why would we have to worry about that, if we don't have to worry about } that when taking the image on film? In fact, we could take care of this } by looking at the image between exposures and correct for z-drift. } However, as you said, that would add to the overall time and exposure. I } was comparing a normal dark field image taken on film at 8 seconds with } acquiring the same image on a "too sensitive" CCD camera by adding up 10 } consecutive .8 second images. Why would the sample drift (in x, y or z) } substantially more in 8+delta seconds than in 8? } } Jim wrote: } } what about the total cost of this additional get-up } } That of course depends on the microscope and there is no general answer. } For example on a LEO 912 I believe the blanker is standard. The } additional cost to use an acquisition scheme like this with our software } is $0 plus perhaps a bit of time to write a small macro. On other } microscopes one might have to add a beam blanker and perhaps a control } mechanism for the beam blanker. But I would guess, that this cost is not } very high. All modern microscopes are computer controller anyway, so it } is most likely just a control command that needs to be sent to the } microscope over a serial port if the beam blanker is installed. Piece of } cake. } } Jim wrote: } } The mind boggles at a through focus series. } } You're right here. But I don't think we were talking about through-focus } series. Incidentally, we do through-focus series on light microscopes } and reconstruction routinely. Takes a few images at different focus (or } for a light microscope: stage) settings. The rest is done off-line. } Takes maybe a couple of minutes for about 20 images of about 1kx1k. I } agree that TEM is different here and much more complicated due to the } complicated Contrast Transfer Function. However, this could in principle } be sorted out. } } Jim wrote: } } Again, I don't doubt that there is now a large place for digital in TEM, } but } its no panacea. } } I also agree with you on that one. But using the additional computer } possibilities of digital imaging might take you further than expected. } } Michael } } } Michael Bode, Ph.D. } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } =================================== } phone: (888) FIND SIS } (303) 234-9270 } fax: (303) 234-9271 } email: mailto:info-at-soft-imaging.com } web: http://www.soft-imaging.com } =================================== } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } Sent: Friday, June 02, 2000 5:15 AM } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } Cc: 'jim-at-proscitech.com.au' } Subject: RE: Film vs Digital } } } The world is full of possible solutions, but are they practical.? } To produce high-resolution, dark-field or any others TEM images that } require } more electrons to form a clear image, Mike Bode would use multiple } digital } exposures. The exposures could be layered and combined into one superior } image. } This image would be made up of more pixel and is formed by more } electrons and } so would be noise-free and hence could be further enlarged then } otherwise } possible. Perhaps. } Beam blanking would largely save the specimen from beam damage and drift } could } be compensated for by matching up the digitals. Great. } How much time is required between exposures to transfer a minimum 10mb } image } per exposure? What would be the total time from focusing to the last } exposure? } What about Z-drift in the interim requiring objective changes and what } about } the total cost of this additional get-up. The mind boggles at a through } focus } series. } When pushing the limits a piece of film seems more effective, cheaper } and fa } ster. } Again, I don't doubt that there is now a large place for digital in TEM, } but } its no panacea. } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com } } On Friday, June 02, 2000 1:24 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] } wrote: } } } } No, it's not really a problem. It's been done with low density } } microscopy all the time. Granted, there are some technical aspects to } be } } overcome, but (and I can only speak for ourselves) we have done that } on } } a number of microscopes. You are of course correct, that 10 images at } } 0.8 seconds take longer than 8 seconds as the image has to be } } transferred, etc. BUT: that's what beam blankers are for. It is pretty } } straightforward to take an image at 0.8 seconds, then blank the beam } } very quickly before taking the next image. That way you get pretty } close } } to the 8 sec total exposure. If there is no beam blanker on the } } microscope, in most cases it can be added. } } } } I am not sure what you mean by "too sensitive". The cameras are } usually } } constructed so that 1 electron from the beam creates between a few } tenth } } to a few counts (these are all statistical data, of course). The well } } width divided by this sensitivity then determines, how many primary } } electrons are needed to fully expose one pixel. For example, if the } well } } width is 50,000 electrons and the sensitivity is 1 count/electron, one } } needs 50,000 primary electrons to fill the well. This translates into } } roughly a 0.4% statistical error. } } } } } From a practical standpoint: You can take images with most cameras } when } } the exposure meter on the microscope reads a couple of seconds without } } overexposing the camera. On the other hand, you can reduce the } intensity } } of the beam until you see single electron events. } } } } The one area where CCD cameras may be too sensitive is diffraction. } The } } normally huge intensity in the transmitted beam often leads to } } saturation. In CCDs this can lead to blooming (the intensity spills } over } } into neighboring pixels). This can be taken care of with special chips } } that have anti-blooming features, but this usually has some other } } drawbacks. Again, this can also be overcome somewhat with multiple } } exposures. Film behaves more civilized here, as it simply stops } } responding to the electrons, but this makes film more or less useless } } for quantitative measurements of diffraction patterns. I have done } } diffraction with CCDs many times and though it does require some } } tweaking, one can get very good results from them. } } } } Michael Bode, Ph.D. } } Soft Imaging System Corp. } } 1675 Carr St., #105N } } Lakewood, CO 80215 } } =================================== } } phone: (888) FIND SIS } } (303) 234-9270 } } fax: (303) 234-9271 } } email: mailto:info-at-soft-imaging.com } } web: http://www.soft-imaging.com } } =================================== } } } } } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } } Sent: Wednesday, May 31, 2000 9:37 PM } } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } } Subject: RE: Film vs Digital } } } } } } } ------------------------------------------------------------------------ } } 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 } } } -----------------------------------------------------------------------. } } } } } } Yes, it could be done "in theory". Somebody would need to figure out } the } } } } software and perhaps modify the hardware. Then we would find that the } } total } } exposure of the specimen to the electron beam maybe a muliple of the } } film's } } exposure. Afterall, an 8 sec film exposure would not amount in digital } } to } } 10x0.8, but we would require considerable time in between exposures. } } Since the } } problems in the discussed circumstances are specimen movement and beam } } damage, } } it seems that taking multiple exposures is a poor option. } } } } Digital cameras are for some situation too sensitive to electron } } exposure. } } Cutting back on electrons is no option since its the electrons that } form } } the } } image in the first instance. } } Much easier in light microscopy . . . insert a neutral density } filter. } } Cheers } } Jim Darley } } ProSciTech Microscopy PLUS } } PO Box 111, Thuringowa QLD 4817 Australia } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } www.proscitech.com } }
****This is a commercial response from a Vendor****
Gary and List,
I am sorry you have had a bad experience with Nikon products. However, you are entitled to your "opinion" and therefore; I am entitled to a response. First, the new Digital Eclipse Camera capable of 12M pixels using a stepper mode is exactly the same technology that the new Zeiss Axiocam and an Olympus model uses which has been very well received by the microscope community.
Next, the successor to E1, E2 (technically, made by Fuji) is the current Nikon D1 (Digital SLR) was NOT intended for microscopes. It CAN go on one, but is limited to Brightfield applications and has no NTSC (video) out for focusing. However, it is by far the standard in Digital Cameras! It has won every Journal Award in it's field and is the definitive choice for Professional Photo-Journalists, including the massive market share Nikon has and the majority of Pulitzer Prize winners. As for Film Scanner products, the Nikon Coolscan line is still the standard in the industry. Especially, high end units with Auto Feeders. All Nikon digital products are not perfect. However, these are technical products that are evolving 6 months at a time and are truly in their infantile state from where they will be in just a few years from now! I would prefer to address your specific problems, but you gave none and just called it "junk". With that mentality, I feel my Pentium 266 computer is "junk", but I do not blame the manufacturer because I own an older model.
As for the Nikon Coolpix line (990), pardon my sarcasm, but where are "all the returns" because we need them to fulfill the 38,000 unit Backorders! This camera is so wildly successful EVERY Dealer (Photo and Microscope) is begging for them. Check out Ebay for example; the cameras are selling for more than List Price. Sound to me like a success if you understand the basics of Economics and Supply and Demand. Is it the perfect microscope camera? No, I don't even think so...but it is the ONLY high resolution (3.34M) digital camera, with live video out, goes on any brand or model microscope for under $1000. Oh, did I forget to mention it also won almost every Magazine Award in its class (not all, it actually received a tie with the Olympus 3030 in one, which is a very nice camera, but does not mount a microscope).
Finally, I have been on this List for many years and have respected the use and rules of this forum. We Vendors for the most part are respectful of the opinions of our users. However, making a "blanket statement" like "stay away from Nikon", does not serve the public well; NOR YOURSELF! If you or any customer has specific problems; we want to know about it; as does any reputable manufacturer for future product improvement. The fact is Nikon has the largest market share in microscopes and professional camera equipment and is climbing very quickly on the digital camera list too. So, in short you are entitled to your opinion, but the rest of the world by far does NOT agree with it............
I personally apologize to any members this correspondence offends. Believe me, my preference is to serve the list and answer technical microscopy questions as I have for many years, but this unfair and libel attack required a response.
Regretfully,
Lawrence Kordon Nikon, Inc. Senior Bioscience Specialist nikon-at-jagunet.com
"Dr. Gary Gaugler" 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 } -----------------------------------------------------------------------. } } At 02:21 PM 6/3/00, 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 } } -----------------------------------------------------------------------. } } } } } } Hello all, } } } } Please be aware that this is a commercial post that may be of interest to } } some of the list members. } } I hope not. } } } Digital light microscope cameras are now available that use a highly touted } } CCD detector: Color or monochrome; fast interline transfer, high sensitivity, } } low noise; 6.7 micron square pixel; 2/3 inch 1.3 Million chip; PCI bus frame } } transfer. This detector is ideal to use in light microscopy applications. } } Now precision technology and fast high capacity cheap computers allow this } } detector to be "Micro Stepped" providing digital images of up to 12 million } } pixels to be captured very quickly and providing file sizes of up to 35 MB in } } 24 bit color images. "Inter Pixel Stepping" technology will allow a } } considerably less expensive approach to digital imaging that can now approach } } film resolving capability in a light microscope. } } } } Hats of to Dr. Jeffree's explanation on optical resolution as it relates to } } light microscopy applications. With high pixel density capable digital } } cameras, now lower magnification, low NA (lower resolving power) objectives } } can be used to acquire matching digital resolution images of wide fields of } } view. } } } } Information on Nikon's Instrument Division "Digital Eclipse" family of } } digital cameras for microscopy including the new DXM1200 digital color camera } } will soon be available on our web site www.nikonusa.com or you can go to } } our Dealer Locator at } } http://www.nikonusa.com/corpinfo/dealers/dealerSearch.cfm and ask your } } local Nikon authorized microscope dealer for further information. } } } } Best Regards, } } } } Stan Schwartz } } Manager, BioSciences Dept. } } Nikon Inc Instrument Division } } 1300 Walt Whitman Rd. } } Melville, NY 11747 } } 631-547-8500 } } 631-547-4033 Fax } } Schwartz-at-nikonincmail.com } } www.nikonusa.com } } As a long term past user of Nikon equipment, it is a tale of sorrow. } Most recently, the digital E1 and E2 are dismal failures. Consumers } are sending back 990's in droves (see rec.photo.marketplace.digital). } I really think that Nikon blew it in regards to digicams. How Nikon can } claim to get a 1.3M pixel imager to produce realistic 12M pixel images } is rather absurd....if not offensive. The most recent D1 is a joke. Albeit, } an expensive one. } } Nikon blew it in the scanner arena and made huge blunders in high end } pro digicams (E1 & E2). Total junk from my personal experiences. I am } not at all prone to spend a dime on any new Nikon digital things. In fact, } I have } dumped my Nikon so-called pro lenses and bodies for Contax. But this } is another story. } } My advice is to be very wary....be very wary of Nikon. I do not use Nikon } cameras anymore and I do not use Nikon microscopes anymore. } But it is your money and your decision. As they say, "Caveat emptor." } } gg } } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ } Modern surfers use PC boards. You can too at } http://photoweb.net } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
It's been some time since I posted news about Project MICRO, MSA's middle school educational outreach program. I'm happy to report that Nestor, wearing his Webmaster hat (yes, he owns more than one), has just posted a substantial revision of the MICRO website (URL below). You'll find new information on several pages and a LOT of new entries in the bibliography. Don't miss the "Cyclops" videos and the new CD-ROMs; the website hotlinks are much expanded also.
Caroline
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
I want to stain Procure-Araldite embedded material using the periodic-acid schiff procedure. One method I have is to hydrolyze in 1N HCl at 60 degrees C for 10 mins, water wash, stain in schiffs, metabisulphate rinses, counterstain, water wash (O'Brien and McCully, 1981). Does anyone use a method routinely which varies from this??
A fellow colleague has bought a JEOL 100s and wants and needs EDS X-ray Analysis. He's mounted his horizintal EDS detector , but can not get sample X-rays from the speciman. Contacting JEOL he found the problem to be a tilt problem. The 10 degree tilt from the normal SEG only tilts 10 degrees and a 30 to 60 degree tilt is necessary. At one time I was told that special sample holders and or double gap pole pieces were availabe. He wishs to find any one with a 100s for parts needed or information which would allow X-ray analysis. Please reply to mgmanders-at-aol.com or the list server.
I reacall some one looking for a desktop incubator. I came a across one on ebay. http://cgi.ebay.com/aw-cgi/eBayISAPI.dll?ViewItem&item=344913999 It is a little pricy by ebay standards. You can find a bunch of incubators by going to www.ebay.com and search for incubators you can do the same a http://www.labx.com
I don't have any connection with anyone involved in this.
Gordon
Gordon Couger gcouger-at-couger.com
Stillwater, OK www.couger.com/gcouger 405 624-2855 GMT -6:00
Toby, I used this technique years ago when I worked for Terry O'Brien at Monash Uni. Depending on the type of tissue you may want to do an aldehyde blockade before you begin the staining procedure. This technique is also in O'Brien and McCully. The blockade removes any aldehyde grouping that will react with the Schiffs reagent. You then generate and stain aldehyde groups during the procedure. Regards JVN
Toby Knight 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 want to stain Procure-Araldite embedded material using the periodic-acid } schiff procedure. One method I have is to hydrolyze in 1N HCl at } 60 degrees C for 10 mins, water wash, stain in schiffs, metabisulphate } rinses, counterstain, water wash (O'Brien and McCully, 1981). Does anyone } use a method routinely which varies from this?? } } thanks in advance, Toby Knight. } } -------------------------------------------------------- } Toby Knight } PhD student } } Department of Horticulture, Viticulture and Oenology } The University of Adelaide } Plant Research Centre } Waite Campus, PMB 1 } Glen Osmond, SA 5064 } } Tel: +61 8 8303 7224 or 8303 6668 } HVO: +61 8 8303 7242 } Fax: +61 8 8303 7116 } Email: tknight-at-waite.adelaide.edu.au } --------------------------------------------------------
-- **************************************************** John V Nailon Operations Manager Centre for Microscopy and Microanalysis The University of Queensland St. Lucia Queensland 4072 Phone: +61-7-3365-4214 Fax: +61-7-3365-4422 WWW: http://www.uq.edu.au/nanoworld/allstaff.html#Nailon ****************************************************
Toby- I've worked with the PAS stain with methyl methacrylate embedded tissue cut at 2 microns. I used a kit from Polyscientific (NY, USA) which supplied me with all of the necessary reagents (minus the ethanol and xylene). The protocol that was supplied with the kit is for paraffin embedded material but I made modifications to the protocol to accommodate methyl methacrylate. If you are interested in the modified protocol, please contact me. I should also mention that I automated this stain to save time and to maximize quality.
Good luck! Michelle Taurino Aventis Pharmaceuticals Bioimaging and Molecular Histology Tel-908-231-3357 Fax-908-231-3962 e-mail: Michelle.Taurino-at-aventis.com
-----Original Message----- } From: Toby Knight [mailto:tknight-at-waite.adelaide.edu.au] Sent: Sunday, June 04, 2000 10:59 PM To: Histonet; Microscopy list
I want to stain Procure-Araldite embedded material using the periodic-acid schiff procedure. One method I have is to hydrolyze in 1N HCl at 60 degrees C for 10 mins, water wash, stain in schiffs, metabisulphate rinses, counterstain, water wash (O'Brien and McCully, 1981). Does anyone use a method routinely which varies from this??
Just like to thank all those who responded to my squeal for help on BS detector resolution. Would like to add that the South African microscopy community is looking into establishing QA procedures and some of the suggestions maybe very helpful in this regard. Thanks, Malc.
-- Dr MP Roberts Phone: [+27](0)46 603 8313 Dept of Geology Fax: [+27](0)46 622 9715 Rhodes University Cell: 083 4060 262 (usually off) 6140 Grahamstown e-mail: malc-at-rock.ru.ac.za SOUTH AFRICA
I've been trying to look at Mycoplasms with SEM. Untill now, the results are not what we hoped for. We've used polycarbonate filters to prepare the Mycoplasms. Another way was preparing the Mycoplasms on agar. We have done this already for enterococcus and we know that this technique works well. But with the Mycoplasms we see almost nothing. It's like the Mycoplasms are IN the Agar. Has anyone have experience with MYCOPLASMS and SEM ???
De Pauw Bart Ghent University Faculty of Veterinary Medicine Morphology Salisburylaan 133 9820 Merelbeke Belgium Phone : 0032(0)9 264.77.19 Fax : 0032(0)9 264.77.90
Toby The procedure you describe is not in fact Periodic Acid-Schiff but the Feulgen reaction, for visualization of DNA in nuclei and mitochondria.
The PAS reaction uses periodic acid or sodium meta-periodate (typically 1% solution, ~10min) to oxidise the vicinal diols of some polysaccharides which are then stained with pararosaniline Schiff's reagent. It is not usually necessary to use metabisulphite in the wash water. The detailed procedure is also described in O'Brien & McCully 1981.
PAS (and Feulgen) may be viewed in a brightfield microscope or by fluorescence microscopy with a "rhodamine" filter set. An alternative fluorescent dye which works well in a fluorescence pseudo Schiff procedure is Lucifer Yellow CH, which needs blue excitation (FITC filter set).
Chris
Date sent: Mon, 5 Jun 2000 12:29:02 +0930 (CST) } From: Toby Knight {tknight-at-waite.adelaide.edu.au} Send reply to: Toby Knight {tknight-at-waite.adelaide.edu.au}
I just came across the following pearl in a 1982 Kodak publication, number PDS 61 "Selecting film from Kodak for photomicrography".
"The (Technical Pan) 2415 film has very low granularity and is capable of very great enlargement. In some cases the resolving power may be beyond that of the microscope image"
How do they know! Answers on a postcard please ....
Chris ===================================================================== DR CHRIS JEFFREE BIOSEM - BIOLOGICAL SCIENCES EM FACILITY UNIVERSITY OF EDINBURGH Daniel Rutherford Building King's Buildings, Mayfield Road EDINBURGH, EH9 3JH, Scotland, UK Tel. #44 131 650 5345 FAX. #44 131 650 6563 Mobile 0410 585 401 email c.jeffree-at-ed.ac.uk SEM / TEM bookings sem-at-ed.ac.uk =====================================================================
The Electron Microscopy Core Facility at Mayo Clinic in Rochester, MN has an opening for a Biomedical EM technologist to support both clinical and research projects. The laboratory offers expertise to collaborative projects that involve transmission and scanning electron microscopy. The laboratory is well equipped and has a history of excellent productivity and adequate funding.
The successful candidate for this position will possess at least a bachelor's degree in Biology with experience in histology and/or electron microscopy. Additional courses or experience in Immunology, Cell Biology, and Digital Imaging is desirable. Operating knowledge of transmission and scanning electron microscopes is preferred. The applicant must have excellent communicative skills and the ability to work well with a variety of personalities.
The EM Technologist interacts with all laboratory users in order to accomplish specific research and clinical goals with respect to electron microscopy procedures. Duties include: All aspects of specimen preparation for a variety of biomedical samples for TEM and SEM, operation of TEM and SEM, darkroom developing and printing, digital image capture, and reporting. The technologist will also perform advanced research procedures including immunoelectron microscopy, x-ray microanalysis, and microwave processing.
Mayo offers a competitive salary and benefits package. If interested, please submit a cover letter and resume referencing job posting #00-0002365 to:
Jill Kelly Mayo Medical Center Human Resources-OE 1 Rochester, MN 55905 Fax: 507-284-1445 Email: kelly.jill-at-mayo.edu
Jon Charlesworth Coordinator Electron Microscopy Core Facility 1426 Gugg X4-3148
'I have two replies and Mike my well have four replies to these.'
I'll try. But I agree with you that we should let this thread die. jim, it seems you're a bit angy at me. If I unintentionally stepped on your toe I apologize. I did not expect that we would agree 100%, as you are selling film and I am involved in digital systems. I just hope that some other readers found some of this useful. I don't consider myself a "digital fanatic". This film vs. digital issue has many more facets, some of which we did not even touch, and which can be just as entertaining.
Jim wrote:
'Mike reinforced and strengthened that argument, finishing with the note that he is glad for film when looking at prints by Ansel Adams.'
Just for the record: I usually do not take a magnifying glass to photos. I mentioned Adams because I like his pictures. If he had taken them with a digital camera I would have liked them just as much. I also like some modern art paintings. That does not mean we should start drawing what we see in the microscopes rather than taking pictures ;-)
Jim wrote:
'Incidentally, from the outset I cited increased "enlargability" to obtain high resolution TEM'
Well, you said 'When great enlargements are required film is superior'. Perhaps I misunderstood. I thought, the term great enlargements referred to the microscope and meant 'small details', which you can take easier with a digital camera (no time delay between seeing something and taking an image, no mechanical vibration due to film movement, etc.). I have said many times before that film may be better if high resolution AND large field of view is required.
Jim wrote:
'in any case: Show Sergey!'
I'm in back-channel correspondence with him.
Michael
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: jim [mailto:jim-at-proscitech.com.au] Sent: Sunday, June 04, 2000 2:34 AM To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com'
I have two replies and Mike my well have four replies to these. 1. There has been a parallel discussion concerning resolution of film versus digital images. Clearly in raw power digital cannot compete since film has multi gigabyte capacity. I added to that thread that what matters is: does digital have enough power and that frequently it would. Mike reinforced and strengthened that argument, finishing with the note that he is glad for film when looking at prints by Ansel Adams. (Ansel Adams until about 30 years ago carted for decades large format cameras through US National Parks, especially Yosemite, producing fantastic landscape photographs and books) Adams' limited edition prints were contacts of 5x7 and 8x10" inch sheet film, probably rated at 400 ISO. The line resolution of such prints much exceeds our eyes' resolution, but still results in superior gradation and detail. TEM film, even when much enlarged has such details too. Why Mike, should we accept 2.5 Mb? That is a splendid file size for SEM, and because of the limited enlargability of light microscopy (concrete ceiling due to wavelengths) it's reasonable, but minimal for light microscopy. TEM can do and deserves better.
Incidentally, from the outset I cited increased "enlargability" to obtain high resolution TEM, as one of films major advantages, since greater depths of field at moderate powers makes high powers through photo enlarging a desirable
technique. The small additional magnification yielded by placing a digital camera lower in the column does not compensate. So greater "enlargability" of digitals would be desirable, but is limited by pixel size.
2 The thread was initiated by Sergey. He had problems visualising certain specimens in dark field. The use of his "beaut" digital TEM camera made things worse. I pointed out that the shorter exposure reduced the number of electrons forming the image, hence more noise. I believe that a good part of Mike's case will be settled in his favour when we hear "Eureka" from Sergey's lab. I don't doubt that much more can be done with digital now and that further improvements are on the way. Mike's "solution" may well be possible, but I don't believe its a snap; in any case: Show Sergey! Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Saturday, June 03, 2000 2:58 AM, Mike Bode [SMTP:mb-at-Soft-Imaging.com] wrote: } } Well, let's see: } } Jim wrote: } } Mike Bode would use multiple digital } exposures. The exposures could be layered and combined into one superior } image. } This image would be made up of more pixel and is formed by more } electrons and } so would be noise-free and hence could be further enlarged then } otherwise } possible. Perhaps. } } No, I did not talk about further enlargements. All I wanted to say is, } that a more noise-free image can be achieved by adding multiple images, } and that this also to some extent helps with drift of the sample during } acquisition. } } Jim wrote: } } How much time is required between exposures to transfer a minimum 10mb } image } per exposure? } } How did you arrive at 10 MB? A 1280x1024 image with 16 bit pixel } information is about 2.5 MB (uncompressed). We acquire about 10 of those } per second and transfer them across the PC bus to the display. Putting } them on them into Memory might add a few tenth of a second. Writing to } HD can be done after all images are acquired. } } Jim wrote: } } What would be the total time from focusing to the last exposure? } What about Z-drift in the interim requiring objective changes } } Why would we have to worry about that, if we don't have to worry about } that when taking the image on film? In fact, we could take care of this } by looking at the image between exposures and correct for z-drift. } However, as you said, that would add to the overall time and exposure. I } was comparing a normal dark field image taken on film at 8 seconds with } acquiring the same image on a "too sensitive" CCD camera by adding up 10 } consecutive .8 second images. Why would the sample drift (in x, y or z) } substantially more in 8+delta seconds than in 8? } } Jim wrote: } } what about the total cost of this additional get-up } } That of course depends on the microscope and there is no general answer. } For example on a LEO 912 I believe the blanker is standard. The } additional cost to use an acquisition scheme like this with our software } is $0 plus perhaps a bit of time to write a small macro. On other } microscopes one might have to add a beam blanker and perhaps a control } mechanism for the beam blanker. But I would guess, that this cost is not } very high. All modern microscopes are computer controller anyway, so it } is most likely just a control command that needs to be sent to the } microscope over a serial port if the beam blanker is installed. Piece of } cake. } } Jim wrote: } } The mind boggles at a through focus series. } } You're right here. But I don't think we were talking about through-focus } series. Incidentally, we do through-focus series on light microscopes } and reconstruction routinely. Takes a few images at different focus (or } for a light microscope: stage) settings. The rest is done off-line. } Takes maybe a couple of minutes for about 20 images of about 1kx1k. I } agree that TEM is different here and much more complicated due to the } complicated Contrast Transfer Function. However, this could in principle } be sorted out. } } Jim wrote: } } Again, I don't doubt that there is now a large place for digital in TEM, } but } its no panacea. } } I also agree with you on that one. But using the additional computer } possibilities of digital imaging might take you further than expected. } } Michael } } } Michael Bode, Ph.D. } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } =================================== } phone: (888) FIND SIS } (303) 234-9270 } fax: (303) 234-9271 } email: mailto:info-at-soft-imaging.com } web: http://www.soft-imaging.com } =================================== } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } Sent: Friday, June 02, 2000 5:15 AM } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } Cc: 'jim-at-proscitech.com.au' } Subject: RE: Film vs Digital } } } The world is full of possible solutions, but are they practical.? } To produce high-resolution, dark-field or any others TEM images that } require } more electrons to form a clear image, Mike Bode would use multiple } digital } exposures. The exposures could be layered and combined into one superior } image. } This image would be made up of more pixel and is formed by more } electrons and } so would be noise-free and hence could be further enlarged then } otherwise } possible. Perhaps. } Beam blanking would largely save the specimen from beam damage and drift } could } be compensated for by matching up the digitals. Great. } How much time is required between exposures to transfer a minimum 10mb } image } per exposure? What would be the total time from focusing to the last } exposure? } What about Z-drift in the interim requiring objective changes and what } about } the total cost of this additional get-up. The mind boggles at a through } focus } series. } When pushing the limits a piece of film seems more effective, cheaper } and fa } ster. } Again, I don't doubt that there is now a large place for digital in TEM, } but } its no panacea. } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com } } On Friday, June 02, 2000 1:24 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] } wrote: } } } } No, it's not really a problem. It's been done with low density } } microscopy all the time. Granted, there are some technical aspects to } be } } overcome, but (and I can only speak for ourselves) we have done that } on } } a number of microscopes. You are of course correct, that 10 images at } } 0.8 seconds take longer than 8 seconds as the image has to be } } transferred, etc. BUT: that's what beam blankers are for. It is pretty } } straightforward to take an image at 0.8 seconds, then blank the beam } } very quickly before taking the next image. That way you get pretty } close } } to the 8 sec total exposure. If there is no beam blanker on the } } microscope, in most cases it can be added. } } } } I am not sure what you mean by "too sensitive". The cameras are } usually } } constructed so that 1 electron from the beam creates between a few } tenth } } to a few counts (these are all statistical data, of course). The well } } width divided by this sensitivity then determines, how many primary } } electrons are needed to fully expose one pixel. For example, if the } well } } width is 50,000 electrons and the sensitivity is 1 count/electron, one } } needs 50,000 primary electrons to fill the well. This translates into } } roughly a 0.4% statistical error. } } } } } From a practical standpoint: You can take images with most cameras } when } } the exposure meter on the microscope reads a couple of seconds without } } overexposing the camera. On the other hand, you can reduce the } intensity } } of the beam until you see single electron events. } } } } The one area where CCD cameras may be too sensitive is diffraction. } The } } normally huge intensity in the transmitted beam often leads to } } saturation. In CCDs this can lead to blooming (the intensity spills } over } } into neighboring pixels). This can be taken care of with special chips } } that have anti-blooming features, but this usually has some other } } drawbacks. Again, this can also be overcome somewhat with multiple } } exposures. Film behaves more civilized here, as it simply stops } } responding to the electrons, but this makes film more or less useless } } for quantitative measurements of diffraction patterns. I have done } } diffraction with CCDs many times and though it does require some } } tweaking, one can get very good results from them. } } } } Michael Bode, Ph.D. } } Soft Imaging System Corp. } } 1675 Carr St., #105N } } Lakewood, CO 80215 } } =================================== } } phone: (888) FIND SIS } } (303) 234-9270 } } fax: (303) 234-9271 } } email: mailto:info-at-soft-imaging.com } } web: http://www.soft-imaging.com } } =================================== } } } } } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } } Sent: Wednesday, May 31, 2000 9:37 PM } } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } } Subject: RE: Film vs Digital } } } } } } } ------------------------------------------------------------------------ } } 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 } } } -----------------------------------------------------------------------. } } } } } } Yes, it could be done "in theory". Somebody would need to figure out } the } } } } software and perhaps modify the hardware. Then we would find that the } } total } } exposure of the specimen to the electron beam maybe a muliple of the } } film's } } exposure. Afterall, an 8 sec film exposure would not amount in digital } } to } } 10x0.8, but we would require considerable time in between exposures. } } Since the } } problems in the discussed circumstances are specimen movement and beam } } damage, } } it seems that taking multiple exposures is a poor option. } } } } Digital cameras are for some situation too sensitive to electron } } exposure. } } Cutting back on electrons is no option since its the electrons that } form } } the } } image in the first instance. } } Much easier in light microscopy . . . insert a neutral density } filter. } } Cheers } } Jim Darley } } ProSciTech Microscopy PLUS } } PO Box 111, Thuringowa QLD 4817 Australia } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } www.proscitech.com } }
Jonathon Low kV is preferred for some biological applications chiefly because biologists experience difficulty getting enough contrast out of their specimens. As Dr Alves de Matos rightly says, low kV is more damaging to specimens than high kV, because the penetrating power of low-kV electrons is lower and therefore more beam energy is lost to the specimen. (We think of this exactly the other way round for SEM, but that's a story for another day!).
The main reason for the apparent shift in biologist's attitudes is that many are now less concerned with cell and tissue-level ultrastructure and increasingly interested in imaging macromolecules. 200kV or 300kV machines are preferred for cryo- microscopy of macromolecules in vitrified ice because the resolution is a lot better than at 80 or 120kV, and also because 200kV causes less specimen damage.
However, the cost of resolution either from higher kVs or the shorter working distance objective lenses favoured by materials scientists is poorer image contrast. Philips therefore supply "Biotwin" versions of some of their TEM range which use a long focal length objective which trading a little resolution for a little more contrast. This optical configuration is coincidentally very good for thickish specimens such as ultrathin biological sections where the ultimate resolution of the machine is pretty much academic anyway.
Images of thick (#100nm) specimens such as FIB sections of microelectronic devices seem crisper and more contrasty in our CM120 Biotwin than in a 200KV machine. Chris
} From: "A.P.Alves de Matos" {apmatos-at-ip.pt} To: {Microscopy-at-sparc5.microscopy.com}
Colleagues....
Some of you have misinterpreted my earlier message. Please feel free to carry on the discussion on film/digitization etc..
I only asked that the "tangental thread" centered about complaints on a specific product be taken off-line before any fire and brimstone starts up between a manufacturer and clients. This is not the forum for that type of feedback.
The previous dicussion should certainly continue as long as necessary.
Nestor Your Friendly Neighborhood SysOp
} } Nestor, } } It would be very unfortunate to squelch this important and central debate } concerning not only the corresponding resolution comparisons but also } differances in performance due to noise and greyscale depth. New methods of } enhansing resolution need to be critiqued as well. } } Dave Barnard } } Wadsworth Center HVEM } NYS Dept. Health } Albany NY
We are looking for the "T-tool" - a kind of fixture for TEM sample polishing. It is from T&T group in USA. We need to know the email address or telephone number of T&T group.
Any information about the T&T group or the T-tool will be highly appreciated!
We are looking to purchase a used JEOL 2010 TEM (LaB6, not FE) in good working condition. I would appreciate replies from anyone who knows if a JEOL 2010 is available or will become available in the next 6 months or so. A STEM unit would increase our interest, but isn't strictly necessary.
Also, we will be looking for a buyer for our JEOL 1200EX STEM which has been under service contract from day 1. It has been a highly reliable microscope and has given years of satisfactory performance.
Dave Joswiak Research Scientist Dept. of Astronomy, 351580 University of Washington Seattle, WA 98195 (206)543-7702 joswiak-at-astro.washington.edu
I am forwarding this request on behalf of Mr. Edoardo Nolfo. If anyone could be of assistance please send the information to him directly or (if you prefer) I will forward it to him.
} I am a student at the University of Oxford and am writing to ask for some } advice. I am currently working on multilayer reflectors in beetles; I plan to } use S.E.M. and T.E.M. to elucidate the fine structure of these reflectors. } Apparently the elytra of the beetles must be sliced very thinly for S.E.M. } analysis. My problem lies in the fact that the elytra are extremely } hard, which } means they cannot be sliced very thinly. Do you have any advice to } offer on how } to soften the elytra for this purpose? Perhaps there is a chemical } that is used } to make samples softer, or a standard technique used to soften very hard } samples. I would be extremely grateful to you if you could help me! Please do } not hesitate to contact me if you require any more information. } } Let me thank you in advance for your help; I look forward to hearing from you } soon.
#################################################################### John J. Bozzola, Ph.D., Director Micro-Imaging and Analysis Center 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 Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
Chris, This is because Kodak use EMs to measure the grain size of their silver halide grains. Tech Pan has a very small grain size (slow film, low ISO/ASA number) and is therefore capable of greater enlargement than film with a larger grains. The larger the grain the faster the film (higher ISO/ASA number). Regards JVN JVN
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 } -----------------------------------------------------------------------. } } I just came across the following pearl in a 1982 Kodak publication, } number PDS 61 "Selecting film from Kodak for photomicrography". } } "The (Technical Pan) 2415 film has very low granularity and is } capable of very great enlargement. In some cases the resolving } power may be beyond that of the microscope image" } } How do they know! } Answers on a postcard please .... } } Chris } ===================================================================== } DR CHRIS JEFFREE } BIOSEM - BIOLOGICAL SCIENCES EM FACILITY } UNIVERSITY OF EDINBURGH } Daniel Rutherford Building } King's Buildings, Mayfield Road } EDINBURGH, EH9 3JH, Scotland, UK } Tel. #44 131 650 5345 } FAX. #44 131 650 6563 } Mobile 0410 585 401 } email c.jeffree-at-ed.ac.uk } SEM / TEM bookings sem-at-ed.ac.uk } =====================================================================
-- **************************************************** John V Nailon Operations Manager Centre for Microscopy and Microanalysis The University of Queensland St. Lucia Queensland 4072 Phone: +61-7-3365-4214 Fax: +61-7-3365-4422 WWW: http://www.uq.edu.au/nanoworld/allstaff.html#Nailon ****************************************************
The "T" tool is a variation of the well known Tripod Polisher¨. We are the manufacturers of the Tripod Polisher¨ and also manufacture the more compact BiPod Polisher. The BiPod polisher offers the smaller size as found on the T-tool while still incorporating our many years of experience in SEM and TEM cross section polishing as well as our expertise in precision machining. I know this doesn't help much in locating the T-tool, but it does offer you the information you need to acquire a tool that will effectively meet your requirements. If you would like additional information on these tools, please feel free to contact me.
David Henriks Vice President TEL: 800-728-2233 (toll free in the USA) South Bay Technology, Inc. +1-949-492-2600 1120 Via Callejon FAX: +1-949-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy.
Message text written by Sha Zhu } ----------------------------------------------------------------------- The Microscopy ListServer -- Sponsor: The Microscopy Society of America
We are looking for the "T-tool" - a kind of fixture for TEM sample polishing. It is from T&T group in USA. We need to know the email address or telephone number of T&T group.
Any information about the T&T group or the T-tool will be highly appreciated!
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Jim wrote:
'I have two replies and Mike my well have four replies to these.'
I'll try. But I agree with you that we should let this thread die. jim, it seems you're a bit angy at me. If I unintentionally stepped on your toe I apologize. I did not expect that we would agree 100%, as you are selling film and I am involved in digital systems. I just hope that some other readers found some of this useful. I don't consider myself a "digital fanatic". This film vs. digital issue has many more facets, some of which we did not even touch, and which can be just as entertaining.
Jim wrote:
'Mike reinforced and strengthened that argument, finishing with the note that he is glad for film when looking at prints by Ansel Adams.'
Just for the record: I usually do not take a magnifying glass to photos. I mentioned Adams because I like his pictures. If he had taken them with a digital camera I would have liked them just as much. I also like some modern art paintings. That does not mean we should start drawing what we see in the microscopes rather than taking pictures ;-)
Jim wrote:
'Incidentally, from the outset I cited increased "enlargability" to obtain high resolution TEM'
Well, you said 'When great enlargements are required film is superior'. Perhaps I misunderstood. I thought, the term great enlargements referred to the microscope and meant 'small details', which you can take easier with a digital camera (no time delay between seeing something and taking an image, no mechanical vibration due to film movement, etc.). I have said many times before that film may be better if high resolution AND large field of view is required.
Jim wrote:
'in any case: Show Sergey!'
I'm in back-channel correspondence with him.
Michael
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: jim [mailto:jim-at-proscitech.com.au] Sent: Sunday, June 04, 2000 2:34 AM To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com'
I have two replies and Mike my well have four replies to these. 1. There has been a parallel discussion concerning resolution of film versus digital images. Clearly in raw power digital cannot compete since film has multi gigabyte capacity. I added to that thread that what matters is: does digital have enough power and that frequently it would. Mike reinforced and strengthened that argument, finishing with the note that he is glad for film when looking at prints by Ansel Adams. (Ansel Adams until about 30 years ago carted for decades large format cameras through US National Parks, especially Yosemite, producing fantastic landscape photographs and books) Adams' limited edition prints were contacts of 5x7 and 8x10" inch sheet film, probably rated at 400 ISO. The line resolution of such prints much exceeds our eyes' resolution, but still results in superior gradation and detail. TEM film, even when much enlarged has such details too. Why Mike, should we accept 2.5 Mb? That is a splendid file size for SEM, and because of the limited enlargability of light microscopy (concrete ceiling due to wavelengths) it's reasonable, but minimal for light microscopy. TEM can do and deserves better.
Incidentally, from the outset I cited increased "enlargability" to obtain high resolution TEM, as one of films major advantages, since greater depths of field at moderate powers makes high powers through photo enlarging a desirable
technique. The small additional magnification yielded by placing a digital camera lower in the column does not compensate. So greater "enlargability" of digitals would be desirable, but is limited by pixel size.
2 The thread was initiated by Sergey. He had problems visualising certain specimens in dark field. The use of his "beaut" digital TEM camera made things worse. I pointed out that the shorter exposure reduced the number of electrons forming the image, hence more noise. I believe that a good part of Mike's case will be settled in his favour when we hear "Eureka" from Sergey's lab. I don't doubt that much more can be done with digital now and that further improvements are on the way. Mike's "solution" may well be possible, but I don't believe its a snap; in any case: Show Sergey! Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Saturday, June 03, 2000 2:58 AM, Mike Bode [SMTP:mb-at-Soft-Imaging.com] wrote: } } Well, let's see: } } Jim wrote: } } Mike Bode would use multiple digital } exposures. The exposures could be layered and combined into one superior } image. } This image would be made up of more pixel and is formed by more } electrons and } so would be noise-free and hence could be further enlarged then } otherwise } possible. Perhaps. } } No, I did not talk about further enlargements. All I wanted to say is, } that a more noise-free image can be achieved by adding multiple images, } and that this also to some extent helps with drift of the sample during } acquisition. } } Jim wrote: } } How much time is required between exposures to transfer a minimum 10mb } image } per exposure? } } How did you arrive at 10 MB? A 1280x1024 image with 16 bit pixel } information is about 2.5 MB (uncompressed). We acquire about 10 of those } per second and transfer them across the PC bus to the display. Putting } them on them into Memory might add a few tenth of a second. Writing to } HD can be done after all images are acquired. } } Jim wrote: } } What would be the total time from focusing to the last exposure? } What about Z-drift in the interim requiring objective changes } } Why would we have to worry about that, if we don't have to worry about } that when taking the image on film? In fact, we could take care of this } by looking at the image between exposures and correct for z-drift. } However, as you said, that would add to the overall time and exposure. I } was comparing a normal dark field image taken on film at 8 seconds with } acquiring the same image on a "too sensitive" CCD camera by adding up 10 } consecutive .8 second images. Why would the sample drift (in x, y or z) } substantially more in 8+delta seconds than in 8? } } Jim wrote: } } what about the total cost of this additional get-up } } That of course depends on the microscope and there is no general answer. } For example on a LEO 912 I believe the blanker is standard. The } additional cost to use an acquisition scheme like this with our software } is $0 plus perhaps a bit of time to write a small macro. On other } microscopes one might have to add a beam blanker and perhaps a control } mechanism for the beam blanker. But I would guess, that this cost is not } very high. All modern microscopes are computer controller anyway, so it } is most likely just a control command that needs to be sent to the } microscope over a serial port if the beam blanker is installed. Piece of } cake. } } Jim wrote: } } The mind boggles at a through focus series. } } You're right here. But I don't think we were talking about through-focus } series. Incidentally, we do through-focus series on light microscopes } and reconstruction routinely. Takes a few images at different focus (or } for a light microscope: stage) settings. The rest is done off-line. } Takes maybe a couple of minutes for about 20 images of about 1kx1k. I } agree that TEM is different here and much more complicated due to the } complicated Contrast Transfer Function. However, this could in principle } be sorted out. } } Jim wrote: } } Again, I don't doubt that there is now a large place for digital in TEM, } but } its no panacea. } } I also agree with you on that one. But using the additional computer } possibilities of digital imaging might take you further than expected. } } Michael } } } Michael Bode, Ph.D. } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } =================================== } phone: (888) FIND SIS } (303) 234-9270 } fax: (303) 234-9271 } email: mailto:info-at-soft-imaging.com } web: http://www.soft-imaging.com } =================================== } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } Sent: Friday, June 02, 2000 5:15 AM } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } Cc: 'jim-at-proscitech.com.au' } Subject: RE: Film vs Digital } } } The world is full of possible solutions, but are they practical.? } To produce high-resolution, dark-field or any others TEM images that } require } more electrons to form a clear image, Mike Bode would use multiple } digital } exposures. The exposures could be layered and combined into one superior } image. } This image would be made up of more pixel and is formed by more } electrons and } so would be noise-free and hence could be further enlarged then } otherwise } possible. Perhaps. } Beam blanking would largely save the specimen from beam damage and drift } could } be compensated for by matching up the digitals. Great. } How much time is required between exposures to transfer a minimum 10mb } image } per exposure? What would be the total time from focusing to the last } exposure? } What about Z-drift in the interim requiring objective changes and what } about } the total cost of this additional get-up. The mind boggles at a through } focus } series. } When pushing the limits a piece of film seems more effective, cheaper } and fa } ster. } Again, I don't doubt that there is now a large place for digital in TEM, } but } its no panacea. } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com } } On Friday, June 02, 2000 1:24 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] } wrote: } } } } No, it's not really a problem. It's been done with low density } } microscopy all the time. Granted, there are some technical aspects to } be } } overcome, but (and I can only speak for ourselves) we have done that } on } } a number of microscopes. You are of course correct, that 10 images at } } 0.8 seconds take longer than 8 seconds as the image has to be } } transferred, etc. BUT: that's what beam blankers are for. It is pretty } } straightforward to take an image at 0.8 seconds, then blank the beam } } very quickly before taking the next image. That way you get pretty } close } } to the 8 sec total exposure. If there is no beam blanker on the } } microscope, in most cases it can be added. } } } } I am not sure what you mean by "too sensitive". The cameras are } usually } } constructed so that 1 electron from the beam creates between a few } tenth } } to a few counts (these are all statistical data, of course). The well } } width divided by this sensitivity then determines, how many primary } } electrons are needed to fully expose one pixel. For example, if the } well } } width is 50,000 electrons and the sensitivity is 1 count/electron, one } } needs 50,000 primary electrons to fill the well. This translates into } } roughly a 0.4% statistical error. } } } } } From a practical standpoint: You can take images with most cameras } when } } the exposure meter on the microscope reads a couple of seconds without } } overexposing the camera. On the other hand, you can reduce the } intensity } } of the beam until you see single electron events. } } } } The one area where CCD cameras may be too sensitive is diffraction. } The } } normally huge intensity in the transmitted beam often leads to } } saturation. In CCDs this can lead to blooming (the intensity spills } over } } into neighboring pixels). This can be taken care of with special chips } } that have anti-blooming features, but this usually has some other } } drawbacks. Again, this can also be overcome somewhat with multiple } } exposures. Film behaves more civilized here, as it simply stops } } responding to the electrons, but this makes film more or less useless } } for quantitative measurements of diffraction patterns. I have done } } diffraction with CCDs many times and though it does require some } } tweaking, one can get very good results from them. } } } } Michael Bode, Ph.D. } } Soft Imaging System Corp. } } 1675 Carr St., #105N } } Lakewood, CO 80215 } } =================================== } } phone: (888) FIND SIS } } (303) 234-9270 } } fax: (303) 234-9271 } } email: mailto:info-at-soft-imaging.com } } web: http://www.soft-imaging.com } } =================================== } } } } } } } } -----Original Message----- } } From: jim [mailto:jim-at-proscitech.com.au] } } Sent: Wednesday, May 31, 2000 9:37 PM } } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } } Subject: RE: Film vs Digital } } } } } } } ------------------------------------------------------------------------ } } 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 } } } -----------------------------------------------------------------------. } } } } } } Yes, it could be done "in theory". Somebody would need to figure out } the } } } } software and perhaps modify the hardware. Then we would find that the } } total } } exposure of the specimen to the electron beam maybe a muliple of the } } film's } } exposure. Afterall, an 8 sec film exposure would not amount in digital } } to } } 10x0.8, but we would require considerable time in between exposures. } } Since the } } problems in the discussed circumstances are specimen movement and beam } } damage, } } it seems that taking multiple exposures is a poor option. } } } } Digital cameras are for some situation too sensitive to electron } } exposure. } } Cutting back on electrons is no option since its the electrons that } form } } the } } image in the first instance. } } Much easier in light microscopy . . . insert a neutral density } filter. } } Cheers } } Jim Darley } } ProSciTech Microscopy PLUS } } PO Box 111, Thuringowa QLD 4817 Australia } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } www.proscitech.com } }
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Jonathon Low kV is preferred for some biological applications chiefly because biologists experience difficulty getting enough contrast out of their specimens. As Dr Alves de Matos rightly says, low kV is more damaging to specimens than high kV, because the penetrating power of low-kV electrons is lower and therefore more beam energy is lost to the specimen. (We think of this exactly the other way round for SEM, but that's a story for another day!).
The main reason for the apparent shift in biologist's attitudes is that many are now less concerned with cell and tissue-level ultrastructure and increasingly interested in imaging macromolecules. 200kV or 300kV machines are preferred for cryo- microscopy of macromolecules in vitrified ice because the resolution is a lot better than at 80 or 120kV, and also because 200kV causes less specimen damage.
However, the cost of resolution either from higher kVs or the shorter working distance objective lenses favoured by materials scientists is poorer image contrast. Philips therefore supply "Biotwin" versions of some of their TEM range which use a long focal length objective which trading a little resolution for a little more contrast. This optical configuration is coincidentally very good for thickish specimens such as ultrathin biological sections where the ultimate resolution of the machine is pretty much academic anyway.
Images of thick (#100nm) specimens such as FIB sections of microelectronic devices seem crisper and more contrasty in our CM120 Biotwin than in a 200KV machine. Chris
} From: "A.P.Alves de Matos" {apmatos-at-ip.pt} To: {Microscopy-at-sparc5.microscopy.com}
Dear all,
1) I am working on oak trees, and I would like to describe the ultrastucture of leaves; also I am looking for a protocol (very detailed) to do it. 2) On oak leaves I would like to localize the superoxyde dismutase(s) enzymes by immunocytochemistry, also I am looking for a protocol (very detailed) to do it.
Thanks in advances. Dr. Didier Le Thiec
-------------------------------------------- Didier Le Thiec I.N.R.A. Centre de Recherches Forestieres Unite d'Ecophysiologie Forestiere Laboratoire de Pollution Atmospherique 54280 Champenoux - France
Following on from the very successful meeting held in 1999, this will review the current state-of-the-art of the UK's various FEG/TEM installations in operation.
The meeting will begin with a special keynote lecture on
Advanced TEM Instrumentation for Solving Critical Problems in Materials Science given by Professor Manfred Rźhle Max-Planck Institute fźr Metallforschung, Stuttgart
This lecture will include some news on the new German FEG/TEM project.
There are now ten FEG/TEM facilities in the UK (some with more than one instrument) and the main programme will consist of a series of invited presentations from each of the sites, reviewing the latest developments in instrumentation and covering specialised techniques such as holography, energy-filtered imaging, spectroscopic imaging, nano-scale analysis, etc.
With JIF, JREI and other special equipment funding now becoming available it is likely that there will be other sophisticated FEG/TEM installations in place in the near future. With this in view the meeting will also include a forward look: what will the next generation of instruments be like?
Speakers:
John Hutchison (University of Oxford)
Key-note Lecture - Professor Manfred RÄhle (Stuttgart) Per Bullough (University of Sheffield) Marin van Heel (Imperial College) John Berriman (MRC, Cambridge) Tony Cullis (University of Sheffield) Ian Jones (University of Birmingham) Rik Brydson (University of Leeds) Jeremy Sloan (University of Oxford) David Cherns (University of Bristol) Stephen McVitie (Glasgow University) Stephen Lloyd (University of Cambridge) John Titchmarsh (University of Oxford) Paul Midgley (University of Cambridge)
The meeting will conclude with the RMS AGM and Presidential Address.
For further details contact: Dr Paul Midgley (EMAG): pam33-at-cam.ac.uk or Dr John Hutchison (RMS): john.hutchison-at-materials.ox.ac.uk
Greetings, Just for the record, Tech Pan is interesting in that you can vary the grain size over an amzingly wide range by choice of developer and exposure conditions. You can technidol LC (I think) and get the teeeny weeeeny grains previously mentioned, or use D19 (I think) and get really high contrast big grains.
Just my few exposed halides, Tobias
} Chris, } This is because Kodak use EMs to measure the grain size of their silver } halide grains. Tech Pan has a very small grain size (slow film, low } ISO/ASA number) and is therefore capable of greater enlargement than } film with a larger grains. The larger the grain the faster the film } (higher ISO/ASA number). } Regards } JVN } JVN } } 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 } } -----------------------------------------------------------------------. } } } } I just came across the following pearl in a 1982 Kodak publication, } } number PDS 61 "Selecting film from Kodak for photomicrography". } } } } "The (Technical Pan) 2415 film has very low granularity and is } } capable of very great enlargement. In some cases the resolving } } power may be beyond that of the microscope image" } } } } How do they know! } } Answers on a postcard please .... } } } } Chris } } ===================================================================== } } DR CHRIS JEFFREE } } BIOSEM - BIOLOGICAL SCIENCES EM FACILITY } } UNIVERSITY OF EDINBURGH } } Daniel Rutherford Building } } King's Buildings, Mayfield Road } } EDINBURGH, EH9 3JH, Scotland, UK } } Tel. #44 131 650 5345 } } FAX. #44 131 650 6563 } } Mobile 0410 585 401 } } email c.jeffree-at-ed.ac.uk } } SEM / TEM bookings sem-at-ed.ac.uk } } ===================================================================== } } -- } **************************************************** } John V Nailon } Operations Manager } Centre for Microscopy and Microanalysis } The University of Queensland } St. Lucia Queensland 4072 } Phone: +61-7-3365-4214 } Fax: +61-7-3365-4422 } WWW: http://www.uq.edu.au/nanoworld/allstaff.html#Nailon } ****************************************************
Two weeks ago I posted a short message about the Ocean Optics spectrometer I had just received for doing Plasma diagnostics. I tried it and have found it very useful for examining a plasma cleaning process and optimizing operating conditions. I purchased a USB 2000 model that plugs into a USB port of a PC for less than $3000. Ocean Optics, 727-733-2447, www.OceanOptics.com
Ronald Vane XEI Scientific NEW WEB SITE! www.SEMCLEAN.com
-----Original Message----- } From: Ronald Vane {RVaneXEI-at-concentric.net} To: MSA listserver {Microscopy-at-sparc5.microscopy.com} ; Dr. Gary Gaugler {gary-at-gaugler.com}
My SEM facility has just been approached, and in the interest in providing what they want for a price based on an average, can I query the SEM community as to what they charge for commercial release of their SEM imagery. I'm looking for $/image, but if you charge on any type of sliding scale, I'd be interested in that too. I believe as well I shouldn't be undercutting commercial facilities, so I'm especially interested in these numbers. Reply to me direct, and I'll respond back to the list with the average, max and min, and no names.
cheerios, =shAf= :o)
{} /\ {\/} /\ {\/} /\ {\/} /\ cogito, ergo zZOooOM /\ {\/} /\ {\/} /\ {\/} /\ {} Michael Shaffer, R.A. - mshaf-at-darkwing.uoregon.edu Geological Science's Electron Probe Facility - University of Oregon http://epmalab.uoregon.edu/
I work for North Carolina State University and we are looking in to using Fluorescence in-situ Hybridization (FISH) for identification of bacteria in biofilm. Do you know of any short courses in FISH here in the USA? I found some out of the country but would prefer to take a course in USA.
Thanks for your time
Tracey L. Daly {tldaly-at-unity.ncsu.edu}
Ron Anderson, IBM, Hopewell Jct., New York, USA. anderron-at-us.ibm.com
IBM Analytical Services; http://www.chips.ibm.com/services/asg
I have an associate who is doing cathodoluminescence studies on materials. He would like to extend his observations into the UV for a limited number of tests on a a few samples. He is presently limited by the optics of the microscope and the fiber optics cable connecting the microscope ocular to the input of the spectrometer. Does anyone have any suggestions for optical microscopes with UV optics that might be available for short term lease or loan that he could consider? Objective magnifications of 5X to 10 X and a 5X or 10X ocular would probably suffice.
Thanks,
Don Marshall
Donald J. Marshall Relion Industries P.O. Box 12 Bedford, MA 01730 Ph: 781-275-4695 FAX: 781-271-0252 email dmrelion-at-world.std.com
Cathodoluminescence, mass spectroscopy, electron beam technology
"A weed is a flower out of place."
(Please note: Do not send email with attachments to this address. Instead, send it to donbarlen-at-aol.com. Thank you.)
The Department of Microscopy and Microanalysis at Abbott Laboratories is recruiting an Electron Microscopist for its Biological Microscopy group. This group provides ultrastructural pathology support for Nonclinical Drug Safety studies, as well as for other biological microscopy projects, such as cell screening, virus identification and counting, and immunolabeling.
Requirements for this position include: a Masters' degree in a biological field, such as cell biology, anatomy, or zoology a thorough understanding of mammalian histology and ultrastructure excellent technical skills including tissue collection at necropsy, tissue and cell processing for TEM, sectioning, staining, operation of electron microscopes and related equipment, darkroom procedures ability to communicate information in technical reports and in oral presentations
Highly desirable, but not essential, are: experience in ultrastructural pathology or toxicologic pathology working knowledge of SEM specimen preparation and instrumentation working knowledge of immunocytochemistry, in situ hybridization, and other labeling methods at light or electron microscopic level familiarity with Good Laboratory Practices
We are looking for a team player with outstanding interpersonal skills and the ability to adjust readily to rapidly changing priorities and shifting deadlines. The ability to communicate clearly, both verbally and in writing, is essential. Careful attention to detail and accuracy are required.
The Department of Microscopy and Microanalysis provides corporate-wide support in Biological and Materials Microscopy to all divisions of Abbott Laboratories. The facility houses two TEMs (a Philips CM12 STEM and a LEO 910), two SEMs (a Philips XL30-FEG and AMRAY 1830i), three EDXS systems, a BioRad confocal scanning laser microscope, several fluorescent and light microscopes (polarized light, DIC), and a Quantimet Image Analysis system. We also have an Arcturus PixCell laser capture microdissection system and a Becton Dickinson FACSCalibur flow cytometer. Microtomes include Reichert Ultracut E and S ultramicrotomes, RMC 6000 XL cryoultrotome, Microm histological microtome, and Microm HM500 cryostat.
Please send letters of application and resumes to:
Jane A. Fagerland, Ph.D. Abbott Laboratories D45M/AP31 200 Abbott Park RD. Abbott Park IL 60064-6202
Dear Microscopists, I'm just curious if something is wrong on my end or I'm unsuscribed against my will. I did not receive any posting during the past two-three weeks, and this is impossible in view of previously received 10-20 postings/day. Is there any explanation? Kris Dr. Kristof Kovacs Associate Professor President, Hungarian Society for Microscopy Phone: +36-(88)-421-684 Fax: +36-(88)-328-643 Mailing Address: University of Veszprem, P.O.Box 158, Veszprem H-8201 Hungary
Dear all, I'm currently having a few problems preparing good cross-section specimens of ceramic thin films on LaAlO3 or NdGaO3 substrates.
I can cut them okay, but thinning them often results in cracking as the thickness goes below 100 microns. I have a Gatan model 623 disk grinder and currently have access to SiC abrasive papers to grit sizes of 1500 or 2000, and also have the Gatan diamond polishing disks and specimen lapping kit.
The problems are as follows: If we use the 1500 or 2000 grit SiC, we can often get the samples thin enough, but with poor surface finish which needs to be improved with dimpling (on both sides), we also risk cracking the sample as SiC papers often contain imperfections.
Using the Gatan specimen lapping kit, I find that as the disk grinder is altered to grind off a further 10 microns, the edge of the sample often catches on the diamond disk and tears some of the diamond coating off, leaving a lump on the surface which then risks catching the specimen and cracking it.
I have one possible alternative approach which I used in Sweden last year (with SiAlON ceramics) involving attaching the sample to a glass slide and polishing it using diamond spray on non-absorbent paper. I may consider trying this with these new materials.
Other ideas would be welcome, however, as would suggestions of how to improve my present method. Please note that I do not have the budget to buy any expensive new polishing equipment (such as a tripod polisher, for instance), so the most welcome suggestions would be those that involve improvements to current techniques, use of different types of polishing consumables, etc..
I hope to hear several suggestions, both from users and from the companies who producing polishing and lapping equipment. Why not post them with the list so we can all benefit from the sharing of experience?
Best wishes
===== Ian MacLaren Beijing Laboratory of Electron Microscopy Chinese Academy of Sciences, P.O. Box 2724 100080 Beijing China General Email: ian.maclaren-at-physics.org Work (esp. large attachments): maclaren-at-image.blem.ac.cn
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Since I am usually more involved in material analysis I am a little lost on the biological field. But now I am in the need of a good review article concerning the application of (analytical) TEM to biological materials. I am not looking for sophisticated latest developements but rather for basic applications with some examples, if possible including some examples of analytical TEM.
Hope to get some input :)
Petra
-------------------------------------------------------------- Dr. Petra Wahlbring Centre de Recherche Public - Gabriel Lippmann Laboratoire d'Analyse des Materiaux (LAM) 162a, av. de la Faiencerie L-1511 Luxembourg tel. +352-466644-402 fax +352-466644-400 e-mail: petra.wahlbring-at-crpgl.lu Visit our WWW site! http://www.crpgl.lu/~wahlbrin
ONLY 8 WEEKS TO GO UNTIL ICHC 2000 (3-8 Sept. York University, UK)
Abstracts are still welcome.
Register by the week or combinations of days.
For more details follow the links to the ICHC 2000 web-site below.
Best wishes
Gary Coulton Dr. Gary Coulton Molecular Pathology Division of Biomedical Sciences Imperial College School of Medicine The Sir Alexander Fleming Building South Kensington London SW7 2AZ
------------------------------------- Announcing the 11th International Congress of Histochemistry and Cytochemistry (ICHC 2000)
"Cell Biology and Imaging Tools for the New Century"
September 3-8, 2000, York, United Kingdom
ICHC 2000 comprises 27 symposia addressing latest developments and applications of histochemistry and cytochemistry in the life sciences including medicine.
Many leading experts to speak
8 weeks to go! Register for the week or by the day!
For further details go to http://www.med.ic.ac.uk/external/ichc_2000
I was happy to leave things at that, but there are a couple of things that Mike has raised that need an answer: Your actual quote concerning Ansel Adams was "Having said that and looking at a print of Ansel Adams, I am glad there is film, though!!" This is rather at odds with the new proclamation } " I mentioned Adams because I like his pictures. If he had taken them with } a digital camera I would have liked them just as much. Lots of people have made excellent compositions and many have make technically superior prints. Adam's has excelled by consistently winning on both account. Its a bit strange, but Mike I'll remind you why you his prints: It is not just Adam's great compositions but also the incredible sharpness and tonal range, which is only possible by huge data density and impossible with a 2.5mb enlarged print. Digital not only has problems with occasional great demands on print magnification (pixel size limitations) and lack of electron density (too short exposures), but because TEM negs, like Adam's prints, are capable of a terrific tonal range and resolution. An image appears better if theoretical data minima are exceeded.
I have been a stickler for showing possible conflict of interest. I realize now that I should have made a disclaimer along the way. I did not, simply because it never occurred to me that I could have a real or imagined gain. My aspirations are different, but since we are all subject to self-delusion I'll add the reality: We have very little mark-up on film and cannot export beyond New Zealand. For Australian research institutions money has been very tight and equipment sales have been low for some years. I doubt that at our exchange rate any digital TEM cameras will be installed here during the next couple of years at least. I think that my special friend Chuck is more likely to retain more film sales than ProSciTech may have lost - if anybody was influenced either way.
The word belief is fitting for all of this. Whatever the arguments, I expect people retain their "film versus digital" beliefs. Mike, we can lead a horse to water, but just try to make it float on its back. When you do, send a picture and I won't care if its digital. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Tuesday, June 06, 2000 1:10 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] wrote: } Jim wrote: } } 'I have two replies and Mike my well have four replies to these.' } } I'll try. But I agree with you that we should let this thread die. jim, } it seems you're a bit angy at me. If I unintentionally stepped on your } toe I apologize. I did not expect that we would agree 100%, as you are } selling film and I am involved in digital systems. I just hope that some } other readers found some of this useful. I don't consider myself a } "digital fanatic". This film vs. digital issue has many more facets, } some of which we did not even touch, and which can be just as } entertaining. } } Jim wrote: } } 'Mike reinforced and strengthened that argument, finishing with the note } that he is glad for film when looking at prints by Ansel Adams.' } } Just for the record: I usually do not take a magnifying glass to photos. } I mentioned Adams because I like his pictures. If he had taken them with } a digital camera I would have liked them just as much. I also like some } modern art paintings. That does not mean we should start drawing what we } see in the microscopes rather than taking pictures ;-) } } Jim wrote: } } 'Incidentally, from the outset I cited increased "enlargability" to } obtain high } resolution TEM' } } Well, you said 'When great enlargements are required film is superior'. } Perhaps I misunderstood. I thought, the term great enlargements referred } to the microscope and meant 'small details', which you can take easier } with a digital camera (no time delay between seeing something and taking } an image, no mechanical vibration due to film movement, etc.). I have } said many times before that film may be better if high resolution AND } large field of view is required. } } Jim wrote: } } 'in any case: Show Sergey!' } } I'm in back-channel correspondence with him. } } } } Michael } } Michael Bode, Ph.D. } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } =================================== } phone: (888) FIND SIS } (303) 234-9270 } fax: (303) 234-9271 } email: mailto:info-at-soft-imaging.com } web: http://www.soft-imaging.com } =================================== } } } } -----Original Message----- } From: jim [mailto:jim-at-proscitech.com.au] } Sent: Sunday, June 04, 2000 2:34 AM } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } Subject: RE: Film vs Digital } } } ------------------------------------------------------------------------ } 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 have two replies and Mike my well have four replies to these. } 1. There has been a parallel discussion concerning resolution of } film versus } digital images. Clearly in raw power digital cannot compete since film } has } multi gigabyte capacity. I added to that thread that what matters is: } does } digital have enough power and that frequently it would. Mike reinforced } and } strengthened that argument, finishing with the note that he is glad for } film } when looking at prints by Ansel Adams. } (Ansel Adams until about 30 years ago carted for decades large format } cameras } through US National Parks, especially Yosemite, producing fantastic } landscape } photographs and books) } Adams' limited edition prints were contacts of 5x7 and 8x10" inch sheet } film, } probably rated at 400 ISO. The line resolution of such prints much } exceeds our } eyes' resolution, but still results in superior gradation and detail. } TEM film, } even when much enlarged has such details too. } Why Mike, should we accept 2.5 Mb? That is a splendid file size for SEM, } and } because of the limited enlargability of light microscopy (concrete } ceiling due } to wavelengths) it's reasonable, but minimal for light microscopy. TEM } can do } and deserves better. } } Incidentally, from the outset I cited increased "enlargability" to } obtain high } resolution TEM, as one of films major advantages, since greater depths } of field } at moderate powers makes high powers through photo enlarging a desirable } } technique. The small additional magnification yielded by placing a } digital } camera lower in the column does not compensate. So greater } "enlargability" of } digitals would be desirable, but is limited by pixel size. } } 2 The thread was initiated by Sergey. He had problems visualising } certain } specimens in dark field. The use of his "beaut" digital TEM camera made } things } worse. I pointed out that the shorter exposure reduced the number of } electrons } forming the image, hence more noise. I believe that a good part of } Mike's case } will be settled in his favour when we hear "Eureka" from Sergey's lab. I } don't } doubt that much more can be done with digital now and that further } improvements } are on the way. Mike's "solution" may well be possible, but I don't } believe its } a snap; in any case: Show Sergey! } Cheers } Jim Darley } ProSciTech Microscopy PLUS } PO Box 111, Thuringowa QLD 4817 Australia } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } Great microscopy catalogue, 500 Links, MSDS, User Notes } www.proscitech.com } } On Saturday, June 03, 2000 2:58 AM, Mike Bode [SMTP:mb-at-Soft-Imaging.com] } wrote: } } } } Well, let's see: } } } } Jim wrote: } } } } Mike Bode would use multiple digital } } exposures. The exposures could be layered and combined into one } superior } } image. } } This image would be made up of more pixel and is formed by more } } electrons and } } so would be noise-free and hence could be further enlarged then } } otherwise } } possible. Perhaps. } } } } No, I did not talk about further enlargements. All I wanted to say is, } } that a more noise-free image can be achieved by adding multiple } images, } } and that this also to some extent helps with drift of the sample } during } } acquisition. } } } } Jim wrote: } } } } How much time is required between exposures to transfer a minimum 10mb } } image } } per exposure? } } } } How did you arrive at 10 MB? A 1280x1024 image with 16 bit pixel } } information is about 2.5 MB (uncompressed). We acquire about 10 of } those } } per second and transfer them across the PC bus to the display. Putting } } them on them into Memory might add a few tenth of a second. Writing to } } HD can be done after all images are acquired. } } } } Jim wrote: } } } } What would be the total time from focusing to the last exposure? } } What about Z-drift in the interim requiring objective changes } } } } Why would we have to worry about that, if we don't have to worry about } } that when taking the image on film? In fact, we could take care of } this } } by looking at the image between exposures and correct for z-drift. } } However, as you said, that would add to the overall time and exposure. } I } } was comparing a normal dark field image taken on film at 8 seconds } with } } acquiring the same image on a "too sensitive" CCD camera by adding up } 10 } } consecutive .8 second images. Why would the sample drift (in x, y or } z) } } substantially more in 8+delta seconds than in 8? } } } } Jim wrote: } } } } what about the total cost of this additional get-up } } } } That of course depends on the microscope and there is no general } answer. } } For example on a LEO 912 I believe the blanker is standard. The } } additional cost to use an acquisition scheme like this with our } software } } is $0 plus perhaps a bit of time to write a small macro. On other } } microscopes one might have to add a beam blanker and perhaps a control } } mechanism for the beam blanker. But I would guess, that this cost is } not } } very high. All modern microscopes are computer controller anyway, so } it } } is most likely just a control command that needs to be sent to the } } microscope over a serial port if the beam blanker is installed. Piece } of } } cake. } } } } Jim wrote: } } } } The mind boggles at a through focus series. } } } } You're right here. But I don't think we were talking about } through-focus } } series. Incidentally, we do through-focus series on light microscopes } } and reconstruction routinely. Takes a few images at different focus } (or } } for a light microscope: stage) settings. The rest is done off-line. } } Takes maybe a couple of minutes for about 20 images of about 1kx1k. I } } agree that TEM is different here and much more complicated due to the } } complicated Contrast Transfer Function. However, this could in } principle } } be sorted out. } } } } Jim wrote: } } } } Again, I don't doubt that there is now a large place for digital in } TEM, } } but } } its no panacea. } } } } I also agree with you on that one. But using the additional computer } } possibilities of digital imaging might take you further than expected. } } } } Michael } } } } } } Michael Bode, Ph.D. } } Soft Imaging System Corp. } } 1675 Carr St., #105N } } Lakewood, CO 80215 } } =================================== } } phone: (888) FIND SIS } } (303) 234-9270 } } fax: (303) 234-9271 } } email: mailto:info-at-soft-imaging.com } } web: http://www.soft-imaging.com } } =================================== } } } } } } } } -----Original Message----- } } } From: jim [mailto:jim-at-proscitech.com.au] } } Sent: Friday, June 02, 2000 5:15 AM } } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } } Cc: 'jim-at-proscitech.com.au' } } Subject: RE: Film vs Digital } } } } } } The world is full of possible solutions, but are they practical.? } } To produce high-resolution, dark-field or any others TEM images that } } require } } more electrons to form a clear image, Mike Bode would use multiple } } digital } } exposures. The exposures could be layered and combined into one } superior } } image. } } This image would be made up of more pixel and is formed by more } } electrons and } } so would be noise-free and hence could be further enlarged then } } otherwise } } possible. Perhaps. } } Beam blanking would largely save the specimen from beam damage and } drift } } could } } be compensated for by matching up the digitals. Great. } } How much time is required between exposures to transfer a minimum 10mb } } image } } per exposure? What would be the total time from focusing to the last } } exposure? } } What about Z-drift in the interim requiring objective changes and what } } about } } the total cost of this additional get-up. The mind boggles at a } through } } focus } } series. } } When pushing the limits a piece of film seems more effective, cheaper } } and fa } } ster. } } Again, I don't doubt that there is now a large place for digital in } TEM, } } but } } its no panacea. } } Cheers } } Jim Darley } } ProSciTech Microscopy PLUS } } PO Box 111, Thuringowa QLD 4817 Australia } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } www.proscitech.com } } } } On Friday, June 02, 2000 1:24 AM, Mike Bode [SMTP:mb-at-soft-imaging.com] } } wrote: } } } } } } No, it's not really a problem. It's been done with low density } } } microscopy all the time. Granted, there are some technical aspects } to } } be } } } overcome, but (and I can only speak for ourselves) we have done that } } on } } } a number of microscopes. You are of course correct, that 10 images } at } } } 0.8 seconds take longer than 8 seconds as the image has to be } } } transferred, etc. BUT: that's what beam blankers are for. It is } pretty } } } straightforward to take an image at 0.8 seconds, then blank the beam } } } very quickly before taking the next image. That way you get pretty } } close } } } to the 8 sec total exposure. If there is no beam blanker on the } } } microscope, in most cases it can be added. } } } } } } I am not sure what you mean by "too sensitive". The cameras are } } usually } } } constructed so that 1 electron from the beam creates between a few } } tenth } } } to a few counts (these are all statistical data, of course). The } well } } } width divided by this sensitivity then determines, how many primary } } } electrons are needed to fully expose one pixel. For example, if the } } well } } } width is 50,000 electrons and the sensitivity is 1 count/electron, } one } } } needs 50,000 primary electrons to fill the well. This translates } into } } } roughly a 0.4% statistical error. } } } } } } } From a practical standpoint: You can take images with most cameras } } when } } } the exposure meter on the microscope reads a couple of seconds } without } } } overexposing the camera. On the other hand, you can reduce the } } intensity } } } of the beam until you see single electron events. } } } } } } The one area where CCD cameras may be too sensitive is diffraction. } } The } } } normally huge intensity in the transmitted beam often leads to } } } saturation. In CCDs this can lead to blooming (the intensity spills } } over } } } into neighboring pixels). This can be taken care of with special } chips } } } that have anti-blooming features, but this usually has some other } } } drawbacks. Again, this can also be overcome somewhat with multiple } } } exposures. Film behaves more civilized here, as it simply stops } } } responding to the electrons, but this makes film more or less } useless } } } for quantitative measurements of diffraction patterns. I have done } } } diffraction with CCDs many times and though it does require some } } } tweaking, one can get very good results from them. } } } } } } Michael Bode, Ph.D. } } } Soft Imaging System Corp. } } } 1675 Carr St., #105N } } } Lakewood, CO 80215 } } } =================================== } } } phone: (888) FIND SIS } } } (303) 234-9270 } } } fax: (303) 234-9271 } } } email: mailto:info-at-soft-imaging.com } } } web: http://www.soft-imaging.com } } } =================================== } } } } } } } } } } } } -----Original Message----- } } } From: jim [mailto:jim-at-proscitech.com.au] } } } Sent: Wednesday, May 31, 2000 9:37 PM } } } To: 'Mike Bode'; 'Microscopy-at-MSA.Microscopy.Com' } } } Subject: RE: Film vs Digital } } } } } } } } } } } } ------------------------------------------------------------------------ } } } 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 } } } } } } -----------------------------------------------------------------------. } } } } } } } } } Yes, it could be done "in theory". Somebody would need to figure out } } the } } } } } } software and perhaps modify the hardware. Then we would find that } the } } } total } } } exposure of the specimen to the electron beam maybe a muliple of the } } } film's } } } exposure. Afterall, an 8 sec film exposure would not amount in } digital } } } to } } } 10x0.8, but we would require considerable time in between exposures. } } } Since the } } } problems in the discussed circumstances are specimen movement and } beam } } } damage, } } } it seems that taking multiple exposures is a poor option. } } } } } } Digital cameras are for some situation too sensitive to electron } } } exposure. } } } Cutting back on electrons is no option since its the electrons that } } form } } } the } } } image in the first instance. } } } Much easier in light microscopy . . . insert a neutral density } } filter. } } } Cheers } } } Jim Darley } } } ProSciTech Microscopy PLUS } } } PO Box 111, Thuringowa QLD 4817 Australia } } } Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com } } } Great microscopy catalogue, 500 Links, MSDS, User Notes } } } www.proscitech.com } } }
This sounds like a perfect application of the small angle cleavage technique. You might want to contact Ray Tweston at Univ. of Illinois because I think that we might have successfully prepared one of these while I visited there. At any rate, the technique is very inexpensive and you can prepare sample of superior quality. Get your hands on John McCaffrey and my paper in the MRS TEM sample Prep IV book (vol 480). It has a detailed description on how to do it. South Bay Technology sells the Microcleave kit and you should contact them as well.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center Guys Run Rd. (packages) P. O. Box 11472 (letters) Pittsburgh, PA 15238-0472
Walck-at-PPG.com
(412) 820-8651 (office) (412) 820-8161 (fax)
} -----Original Message----- } From: Ian MacLaren [mailto:maclariz-at-yahoo.co.uk] } Sent: Wednesday, June 07, 2000 3:28 AM } To: Microscopy list } Subject: Ceramic cross-section preparation } } } -------------------------------------------------------------- } ---------- } 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'm currently having a few problems preparing good } cross-section specimens } of ceramic thin films on LaAlO3 or NdGaO3 substrates. } } I can cut them okay, but thinning them often results in } cracking as the } thickness goes below 100 microns. I have a Gatan model 623 } disk grinder and } currently have access to SiC abrasive papers to grit sizes of } 1500 or 2000, } and also have the Gatan diamond polishing disks and specimen } lapping kit. } } The problems are as follows: } If we use the 1500 or 2000 grit SiC, we can often get the } samples thin } enough, but with poor surface finish which needs to be improved with } dimpling (on both sides), we also risk cracking the sample as } SiC papers } often contain imperfections. } } Using the Gatan specimen lapping kit, I find that as the disk } grinder is } altered to grind off a further 10 microns, the edge of the } sample often } catches on the diamond disk and tears some of the diamond } coating off, } leaving a lump on the surface which then risks catching the } specimen and } cracking it. } } I have one possible alternative approach which I used in } Sweden last year } (with SiAlON ceramics) involving attaching the sample to a } glass slide and } polishing it using diamond spray on non-absorbent paper. I } may consider } trying this with these new materials. } } Other ideas would be welcome, however, as would suggestions of how to } improve my present method. Please note that I do not have } the budget to buy } any expensive new polishing equipment (such as a tripod polisher, for } instance), so the most welcome suggestions would be those } that involve } improvements to current techniques, use of different types of } polishing } consumables, etc.. } } I hope to hear several suggestions, both from users and from } the companies } who producing polishing and lapping equipment. Why not post } them with the } list so we can all benefit from the sharing of experience? } } Best wishes } } ===== } Ian MacLaren } Beijing Laboratory of Electron Microscopy } Chinese Academy of Sciences, P.O. Box 2724 } 100080 Beijing } China } General Email: ian.maclaren-at-physics.org } Work (esp. large attachments): maclaren-at-image.blem.ac.cn } } ____________________________________________________________ } Do You Yahoo!? } Get your free -at-yahoo.co.uk address at http://mail.yahoo.co.uk } or your free -at-yahoo.ie address at http://mail.yahoo.ie }
Postdoctoral Position Immediately Available In situ UHV-TEM of Nanoparticle Reactions and Metal Oxidation
Materials Research Laboratory, University of Illinois at Urbana-Champaign and University of Pittsburgh
A postdoctoral position is immediately available in the Materials Research Laboratory at the University of Illinois at Urbana-Champaign in the area of nano-reactions and in situ UHV-TEM. This position is jointly sponsored between Professor Robert Averback (University of Illinois at Urbana-Champaign) and Professor Judith Yang (University of Pittsburgh).
The research project is two-fold: 1. Surface oxidation kinetics of metals and 2. Nanoparticle reactions/sintering. The research project is to combine the unique experimental information obtainable from in situ UHV-TEM and compare with theoretical models of these nanoscale reactions. The position requires a PhD in physical sciences/engineering. Hands-on experience in TEM techniques is highly desirable. The position is open to all qualified candidates and has an anticipated duration of 2 years. If you are interested in this opportunity, please send a resume and names of three references to the address below.
Dr. Judith C. Yang Assistant Professor Dept. of Materials Science & Engineering 848 Benedum Hall University of Pittsburgh Pittsburgh, PA 15261
I can reccomend you the following books: 1. D.C.Joy, A.D.Romig and J.I.Goldstein, "Principles of analytical electron microscopy", Plenum Press, 1989; it contains three chapters dedicated to bilogy: chapter 6, 12 and 13 2. J.J.Hren, J.I.Goldstein and D.C.Joy, "Introduction to AEM", Plenum Press, 1979; it contains chapters dedicated to biology. I hope this helps.
Corneliu Sarbu Dept.MTM KULeuven Belgium
- - - - - - - - - - - - - - Original Message - - - - - - - - - - - - - - ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Since I am usually more involved in material analysis I am a little lost on the biological field. But now I am in the need of a good review article concerning the application of (analytical) TEM to biological materials. I am not looking for sophisticated latest developements but rather for basic applications with some examples, if possible including some examples of analytical TEM.
Hope to get some input :)
Petra
-------------------------------------------------------------- Dr. Petra Wahlbring Centre de Recherche Public - Gabriel Lippmann Laboratoire d'Analyse des Materiaux (LAM) 162a, av. de la Faiencerie L-1511 Luxembourg tel. +352-466644-402 fax +352-466644-400 e-mail: petra.wahlbring-at-crpgl.lu Visit our WWW site! http://www.crpgl.lu/~wahlbrin
- - - - - - - - - - - - End of Original Message - - - - - - - - - - - -
Question: What is the proper procedure for increasing the filament current on a 2010 with a LaB6? How fast can you go up? Should the current be increased at a linear rate or should the rate be tapered off as you reach saturation? The manual suggests 30 sec/graduation while the service techs suggest a much slower rate.
Thanks, Kim ************************************************************** Kim W. Pierson, Ph.D. Dept of Physics & Astronmoy University of Wisconsin-Eau Claire (715) 836-5009 FAX 836-3955
I have a few comments for you here that may help. This is a little touchy as I don't want to knock my competitor's equipment. The Model 623 Disk Grinder that you have is going to cause problems as you describe because of it's design. The way that grinder works is that you use the dial on the top to make your sample extend below the base of the grinder. When you begin polishing, the entire weight of the grinder plus whatever weight you are applying by hand is being transferred to your very thin 3mm diameter sample. In addition to the additional weight, you can tend to apply too much pressure on one side of the sample which would cause the sample to "catch" on the diamond film as you describe.
We manufacture a series of polishing fixtures that are designed to prevent these problems. They are gravity feed fixtures. On these fixtures, the dial gauge at the top simply sets the amount of material that will be fed into the abrasive film. The only weight your sample sees is the weight of the central piston (with some of our fixtures, this weight can be counterbalanced to approach zero). The weight of the rest of the fixture is never transferred to the sample. Also, the sample surface that is being polished is always co-planar with the base of the polishing fixture. This ensures that you cannot apply uneven pressure and that you won't experience the "catching" that you described.
Now the good news. There is one of these fixtures (Model 150) already in use in your facility. There is also the Tripod Polisher¨, Model 920 Lapping & Polishing Machine, Model 850 Wire Saw and Model 650 Diamond Wheel Saw. By separate e-mail,I will give you the details on where these are located and who you should contact to access them.
I also saw Scott Walck's post on the Microcleave technique. I will include a PDF of the paper he was talking about by separate e-mail.
If you have any questions or if I can be of any additional assistance, please contact me directly off-line. I hope this helps.
DISCLAIMER: South Bay Technology produces equipment and supplies as described above and, therefore, has a vested interest in promoting their use.
David Henriks Vice President TEL: 800-728-2233 (toll free in the USA) South Bay Technology, Inc. +1-949-492-2600 1120 Via Callejon FAX: +1-949-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy. Message text written by INTERNET:ian.maclaren-at-physics.org } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear all, I'm currently having a few problems preparing good cross-section specimens of ceramic thin films on LaAlO3 or NdGaO3 substrates.
I can cut them okay, but thinning them often results in cracking as the thickness goes below 100 microns. I have a Gatan model 623 disk grinder and currently have access to SiC abrasive papers to grit sizes of 1500 or 2000,
and also have the Gatan diamond polishing disks and specimen lapping kit.
The problems are as follows: If we use the 1500 or 2000 grit SiC, we can often get the samples thin enough, but with poor surface finish which needs to be improved with dimpling (on both sides), we also risk cracking the sample as SiC papers often contain imperfections.
Using the Gatan specimen lapping kit, I find that as the disk grinder is altered to grind off a further 10 microns, the edge of the sample often catches on the diamond disk and tears some of the diamond coating off, leaving a lump on the surface which then risks catching the specimen and cracking it.
I have one possible alternative approach which I used in Sweden last year (with SiAlON ceramics) involving attaching the sample to a glass slide and polishing it using diamond spray on non-absorbent paper. I may consider trying this with these new materials.
Other ideas would be welcome, however, as would suggestions of how to improve my present method. Please note that I do not have the budget to buy any expensive new polishing equipment (such as a tripod polisher, for instance), so the most welcome suggestions would be those that involve improvements to current techniques, use of different types of polishing consumables, etc..
I hope to hear several suggestions, both from users and from the companies who producing polishing and lapping equipment. Why not post them with the list so we can all benefit from the sharing of experience?
I'm trying to do immunogold studies on a membrane protein in a preparation of plant vesicles (from minus 80ĽC storage). I did an Epon embedding with standard procedure (glut and osmium) to check the ultrastructure and got decent-looking vesicles, but side by side an LR White prep (0.5% glut, 2% paraformaldehyde, without osmium) looks like a disaster, with granular material, vague membranish-like structures and whorls of membranes. Does anyone have any tips for LR White preps of vesicles, or any other ideas for embedding (resins, fixing, dehydration, embedding) to favour the immunogold process.
Thanks,
Mark
******************************************** Mark West, Unidad de Microscopia Electronica, (Electron Microscopy Unit) Instituto de Fisiologia Celular, Universidad Nacional Autonoma de Mexico, 04510 Mexico D.F.
A book that I have found to be very resourceful is Biomedical Electron Microscopy by Arvid B. Maunsbach and Bjorn Afzelius.
Hope this is useful to you as well. Regards- Michelle Taurino Aventis Pharmaceuticals Bioimaging and Molecular Histology Tel-908-231-3357 Fax-908-231-3962 e-mail: Michelle.Taurino-at-aventis.com
-----Original Message----- } From: Petra Wahlbring [mailto:wahlbrin-at-crpgl.lu] Sent: Wednesday, June 07, 2000 3:49 AM To: microscopy-at-sparc5.microscopy.com
Since I am usually more involved in material analysis I am a little lost on the biological field. But now I am in the need of a good review article concerning the application of (analytical) TEM to biological materials. I am not looking for sophisticated latest developements but rather for basic applications with some examples, if possible including some examples of analytical TEM.
Hope to get some input :)
Petra
-------------------------------------------------------------- Dr. Petra Wahlbring Centre de Recherche Public - Gabriel Lippmann Laboratoire d'Analyse des Materiaux (LAM) 162a, av. de la Faiencerie L-1511 Luxembourg tel. +352-466644-402 fax +352-466644-400 e-mail: petra.wahlbring-at-crpgl.lu Visit our WWW site! http://www.crpgl.lu/~wahlbrin
} My SEM facility has just been approached, and in the interest in } providing what they want for a price based on an average, can I query } the SEM community as to what they charge for commercial release of } their SEM imagery. I'm looking for $/image, but if you charge on any } type of sliding scale, I'd be interested in that too. I believe as } well I shouldn't be undercutting commercial facilities, so I'm } especially interested in these numbers. } Reply to me direct, and I'll respond back to the list with the } average, max and min, and no names.
shAf-
Please start with an inquiry to your university administration; you might save yourself a lot of grief. When I was running an interdepartmental lab at U.C. Berkeley, there were very specific university-wide rules that I was required to follow for such usage.
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
We have a 2010 and our procedure is to bring the filament dial (labeled off-10) to position #3. Then we go for coffee. After 20 min. or so we raise it to # 4, after a couple of minutes to #5 etc. until we get to the stop ( in our case we have it at about 7 1/2). Even at this point we have filament drift for quite some time, so , we are not ready to take pictures for another 1/2 hr or so. By the way, we have the bias settings at 7 and 7 which is high. I would be interested in knowing what your settings are.
At 14:19 +0200 3/6/00, Jonathan Barnard wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
My understanding is as follows ....
There are different damage mechnisms at different voltages.
At lower voltages ( {200 kV) the main damage mechanism is, in effect, specimen heating. And, the lower the voltage, the greater the interaction (elastic scattering) with the specimen - so more damage but greater contrast, the main reason users looking at resin sections prefer lower voltages.
Increasing the voltage reduces specimen damage and tends to improve resolution but at the expense of contrast. As the interest in biological TEM moves to molecular biology, resolution is more important. But, at high resolution, phase contrast becomes the dominant factor in producing image constrast and phase constrast can be significantly improved - without loss of resolution - by using highly coherent FEG sources.
At voltages between 300kV and 400 kV, the energy of the electron becomes sufficient to cause "direct" radiation damage - atoms are displaced. At this point, specimen damage rates increase. The particular voltage depends on the amtomic number of your specimen.
Damage rates can also be reduced by cooling. in particular, cooling a specimen to liquid helium temperatures allows significantly greater electron exposure before damage occurs.
So, for optimum imaging of molecular specimens you need 300 kV and a FEG gun plus a liquid helium stage.
Regards, -- Larry Stoter JEOL (UK) Ltd Silver Court, Watchmead, Welwyn Garden City, AL7 1LT, United Kingdom tel: +44-(0)1707-377117, fax: +44-(0)1707-373254, e-mail: larrys-at-jeoleuro.com
For EDS checking you can use specimen from Cu or Al foil so that it will have any angle, including 30-60 degrees. But I am afraid that if there is no spectra at all with 10 degree tilt then a problem is more complicated.
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: "MGMANDERS-at-aol.com"-at-sparc5.microscopy.com } [mailto:"MGMANDERS-at-aol.com"-at-sparc5.microscopy.com] } Sent: Sunday, June 04, 2000 10:41 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: NEED HELP WITH EDS ON A JEOL 100S } } } -------------------------------------------------------------- } ---------- } 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/MicroscopyListser} ver/FAQ.html } } } -------------------------------------------------------------- } ---------. } } } A fellow colleague has bought a JEOL 100s and wants and } needs EDS X-ray } Analysis. He's mounted his horizintal EDS detector , but can } not get sample } X-rays from the speciman. Contacting JEOL he found the } problem to be a tilt } problem. The 10 degree tilt from the normal SEG only tilts } 10 degrees and a } 30 to 60 degree tilt is necessary. At one time I was told } that special } sample holders and or double gap pole pieces were availabe. } He wishs to find } any one with a 100s for parts needed or information which } would allow X-ray } analysis. Please reply to mgmanders-at-aol.com or the list server. } } Mike Manders } } }
We have the opportunity to acquire a TEM, but budget constraints mean that have to consider the cost of acquiring and operating this instrument carefully against alternatives. We plan to use it principally to study the morphology, behavior and size distribution of colloidal gold particles and other types of metal nanoparticles.
Can anyone give any information on what the maintenance, operating and facilities requirements and costs would be for this instrument, or point me to a good source? What renovations would be necessary to accommodate it - darkroom, ancillary equipment, cooling, water and power supplies, darkroom facilities? How much should we budget for supplies and consumables? Since we have no-one currently trained to use it, how long would it take to learn, and how much maintenance (time, supplies, contracts) would it require?
Thanks in advance,
Rick Powell
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Both Zeiss and Leica have long-standing reputations for quartz optics with transmission into the UV (typically, down to about 220nm). You are going to need more than just the objective; the binoc also needs to have a UV transmitting prism. A tip: inquire about microscopes used for microspectrophotometry or in semiconductor applications. From the biological/biomed realm: microscopes used for Fura or Indo studies.
Try Tom Calahan at Zeiss (914-681-7733) and Jan Hinsch at Leica/Allentown (201-236-5905).
Nikon has also been agressive in lens development, but I haven't had direct experience with any UV optics. Call Stan Schwartz (516-547-8529). ... and at Olympus: Reinhard Enders (516-844-5000).
Good hunting! Barbara Foster Microscopy/Microscopy Education 125 Paridon Street, Suitee 102 Springfield, MA 01118
Customized, on-site short courses in all areas of microscopy, sample prep, and image analysis. "Why didn't they teach us that sooner?" Probably because no one called MME!
At 03:27 PM 6/6/00 -0400, donald j marshall wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Suggest you buy a good upright stand which you can upgrade as your needs develop. I strongly suggest a combination of both transmitted and reflected light options.
Your feelings re: investing in the microscope vs. a high end camera at this stage are well founded.
Re: contrast techniques... I think that you might find Hoffman Modulation Contrast a better bet for what you are doing than Phase. Hoffman can be used singly for looking at surfaces (ex: your coatings) or in combination with polarized light. It is a good complement to DIC, which won't work if your powders, etc. respond to Pol.
All of the "big 4" (Olympus, Nikon, Leica, and Zeiss) have good equipment. The issues I would add to your shopping list might include how comfortable you feel operating a specific microscope (like trying on a coat or test-driving a new car) and how supportive your local dealer or representative is.
Your comment re: EM was interesting.... I have been doing a lot of work over the past 6 months with commercial companies who look at many of the applications you cited. Universally, they were astounded at the information they could get from the Light Microscope, quickly and with minimal sample prep. I am currently on assignment teaching a group of very competent EM people about Light Microscopy... and they are equally amazed. All by themselves, they came to the conclusion that they should take a look first with the Light Microscope and then, if necessary, go the SEM. (I added that they should also take a quick look with the stereo first).
By the way, you may be interesting in "Optimizing Light Microscopy", both as a reference for your lab and a text for your students. Details are on our website.
Customized, on-site short courses in all areas of microscopy, sample prep, and image analysis.
"Why didn't they teach us that sooner?" Probably because no one called MME!
At 09:47 AM 6/7/00 +0200, Bemporad, Edoardo wrote:
} } } }
{excerpt}
{fontfamily} {param} Arial {/param} {smaller} I am going to buy one ore two OM for our EM lab (XL30 and CM120), trying to convince myself that one brand is better than the other (quality/price rate included in the evaluation!).
{/smaller} {/fontfamily}
{fontfamily} {param} Arial {/param} {smaller} In our lab we do mainly metallographic analysis, interface studies of wear resistant coatings, and catalytic powders characterization, but I guess that the OM will be used for a wider range of investigation (W/O and O/W emulsions, asbestos, ..) and for didactical scopes too.
{/smaller} {/fontfamily}
{fontfamily} {param} Arial {/param} {smaller} So EPI and DIA illuminations, BF, DF, NIC, phase contrast, pol, I don't think we will need fluorescence; forgotten something?
{/smaller} {/fontfamily}
{fontfamily} {param} Arial {/param} {smaller} What about an inverse microscope? or always better two standard ones? {/smaller} {/fontfamily}
{fontfamily} {param} Arial {/param} {smaller} I have about 35K$ budget to make everybody happy (research group and students), and my position is to prefer spending on the microscope (or microscopes) rather than on a high-level digital camera. I read some threads about it here and I think something like a Nikon coolpix 990 (if it will works! :-) ) will be enough. (other opinions or point of views?)
{/smaller} {/fontfamily}
{fontfamily} {param} Arial {/param} {smaller} I do not have a so deep experience in OM but I was wandering if there are some key feature to keep in mind for an equipment that will be used in a EM lab, considering that where I will not able to go with the OM (depth of field, resolution) I can use our EM!
Most of the lenses should come off, either directly or with the help of a small Allen wrench.
My favorite approach to lens cleaning: "Puff, Huff, and Swirl".... 1. Puff off any debris or rough material which might scratch the lens using either a puffer (available from photographic supply houses) or just "puffing" the dry air from your mouth with your cheeks (easier to demo than explain). 2. Huff on the lens, using the warm, moist air from deep in your lungs, to deposit a fog of moisture on the lens then 3. Quickly and gently, remove the fog with a clean Q-tip (100% cotton ) or lens tissue (NOT Kim-Wipe!), starting at the middle of the lens and continuing in a spiral to the outer edge. Do not use the same area on the swab or tissue more than once; they will collect debris which can scratch the delicate coating. 4. If the dirt is persistent (ex: mascara, fingerprint, oily residue), dip the tip of a cotton swab in a good lens cleaning solution (also available at photo supply houses), shake off any excess, then repeat the "swirl" step.
I recently had a client who had a neat moist towelette made by Uvex. It worked really well, even on oil. I will have to find the contact info, so if you are interested, please email me off line.
Best regards, Barbara Foster Microscopy/Microscopy Education 125 Paridon Street, Suitee 102 Springfield, MA 01118
Customized, on-site short courses in all areas of microscopy, sample prep, and image analysis. "Why didn't they teach us that sooner?" Probably because no one called MME!
At 11:43 PM 6/6/00 -0500, wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
The real question is why should you increase the LaB6 heating current slowly? As I understand it there are two reasons. Firstly if the tip is new, the gun has been serviced or the gun has not been run at 200kV for some time it is important to run up slowly to ensure that you do not get any outgassing from the tip or other gun components that might cause a flashover and possible damage. Secondly if the machine is regularly used at 200kV and the gun is is good condition (normal case) then you want to avoid any thermal shock to the LaB6 tip which could cause damage or misalignment. The same applies to cooling the tip down after use.
In the first case I would certainly take several 10s of minutes while carefully watching the vacuum gauge and the emission current meter (and HT stability if you are connected to an oscilloscope). Exact time would depend on the condition but maybe 30 minutes to heat a new tip and 2 minutes for a normal tip in good condition. I would take a minute to cool down a tip.
In both cases the greatest heating effect is a square of the control position (power is proportional to V^2 or I^2) and this should be taken into acount. I heat to about 3.5 in 25 to 30 seconds then decrease my heating speed as I approach the stop (set at 5 in my case, bias 5.5). The particular setting will depend on the type of LaB6 tip used, the wenhelt to tip distance and how you run the emission. We use 5uA emission with the tip slightly undersaturated as it seems to give good results and reasonable long life at the high mags that we use. For the smallest probes we may desaturate further to reduce the probe size.
I am aware from visitors that we have from other sites that this is quicker than many people but it seems impractical to spend 30 minutes to run up the tip every time you want to change a specimen when we want to look at several in a session. Our tip life des not seem to be any worse than those who take much longer.
Regards, Ron
} ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } Question: } What is the proper procedure for increasing the filament current on a 2010 } with a LaB6? } How fast can you go up? } Should the current be increased at a linear rate or should the rate be } tapered off as you reach saturation? } The manual suggests 30 sec/graduation while the service techs suggest a } much slower rate. } } Thanks, } Kim } ************************************************************** } Kim W. Pierson, Ph.D. } Dept of Physics & Astronmoy } University of Wisconsin-Eau Claire } (715) 836-5009 } FAX 836-3955 } } }
=========================================================================== Mr. Ron Doole e-mail ron.doole-at-materials.ox.ac.uk Department of Materials, phone +44 (0) 1865 273701 University of Oxford, fax +44 (0) 1865 283333 Parks Road. Oxford. OX1 3PH. UK. ============================================================================
LR White won't look as good as Spurr's, but membrane whorls are indicative of insufficient fixation. Os is the better lipid (therefore membrane) fixative, since you cannot use Os, you may need to increase the time and or concentration of GA. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Thursday, June 08, 2000 1:51 AM, Mark West [SMTP:mwest-at-ifcsun1.ifisiol.unam.mx] wrote:
} } Hi, } } I'm trying to do immunogold studies on a membrane protein in a preparation } of plant vesicles (from minus 80oC storage). I did an Epon embedding with } standard procedure (glut and osmium) to check the ultrastructure and got } decent-looking vesicles, but side by side an LR White prep (0.5% glut, 2% } paraformaldehyde, without osmium) looks like a disaster, with granular } material, vague membranish-like structures and whorls of membranes. Does } anyone have any tips for LR White preps of vesicles, or any other ideas } for embedding (resins, fixing, dehydration, embedding) to favour the } immunogold process. } } Thanks, } } Mark } } } } } } } ******************************************** } Mark West, } Unidad de Microscopia Electronica, } (Electron Microscopy Unit) } Instituto de Fisiologia Celular, } Universidad Nacional Autonoma de Mexico, } 04510 Mexico D.F. } } tel (unidad/lab) *(525) 622 5610* } (casa/home) (525) 619 3020 } Fax (525) 616 2282 } ******************************************** }
Hi Kris, same thing happened to me, but re-subscribing took care of the problem. Mailed to listserver, but no respons at all.
} Dear Microscopists, } I'm just curious if something is wrong on my end or I'm unsuscribed against } my will. I did not receive any posting during the past two-three weeks, and } this is impossible in view of previously received 10-20 postings/day. } Is there any explanation? } Kris } Dr. Kristof Kovacs } Associate Professor } President, Hungarian Society for Microscopy } Phone: +36-(88)-421-684 } Fax: +36-(88)-328-643 } Mailing Address: } University of Veszprem, P.O.Box 158, Veszprem } H-8201 Hungary
Yours sincerely
Per Hšrstedt Department of Medical Biosciences Pathology Unit for Electron Microscopy University of UmeŚ S-90187 UmeŚ Sweden
Literature indicates that specimen damage due to heating decreases with increasing accelerating voltage; however, there is a trade off because at higher voltages materials fall victim to knock-on and sputtering damage. A very good overview (with references) of specimen/beam interactions and beam damage can be found on PP 49-55 "Transmission Electron Microscopy" by Williams and Carter 1996 Plenum Press ISBN: 0-306-45342-X
Brenda I. Prenitzer, Ph.D. Member of Technical Staff Cirent Semiconductor (Lucent Technologies) 9333 S. John Young Parkway 6D-Lab Orlando, FL 32819-8612
-----Original Message----- } From: Larry Stoter [mailto:LPS-at-teknesis.demon.co.uk] Sent: Wednesday, June 07, 2000 3:49 PM To: Jonathan Barnard; microscopy-at-sparc5.microscopy.com
At 14:19 +0200 3/6/00, Jonathan Barnard wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
My understanding is as follows ....
There are different damage mechnisms at different voltages.
At lower voltages ( {200 kV) the main damage mechanism is, in effect, specimen heating. And, the lower the voltage, the greater the interaction (elastic scattering) with the specimen - so more damage but greater contrast, the main reason users looking at resin sections prefer lower voltages.
Increasing the voltage reduces specimen damage and tends to improve resolution but at the expense of contrast. As the interest in biological TEM moves to molecular biology, resolution is more important. But, at high resolution, phase contrast becomes the dominant factor in producing image constrast and phase constrast can be significantly improved - without loss of resolution - by using highly coherent FEG sources.
At voltages between 300kV and 400 kV, the energy of the electron becomes sufficient to cause "direct" radiation damage - atoms are displaced. At this point, specimen damage rates increase. The particular voltage depends on the amtomic number of your specimen.
Damage rates can also be reduced by cooling. in particular, cooling a specimen to liquid helium temperatures allows significantly greater electron exposure before damage occurs.
So, for optimum imaging of molecular specimens you need 300 kV and a FEG gun plus a liquid helium stage.
Regards, -- Larry Stoter JEOL (UK) Ltd Silver Court, Watchmead, Welwyn Garden City, AL7 1LT, United Kingdom tel: +44-(0)1707-377117, fax: +44-(0)1707-373254, e-mail: larrys-at-jeoleuro.com
I remember a message some time ago, I think, suggesting a way to flat embed in silicon molds using LRWhite resin. How the top was sealed escapes me. Could anyone with a better filing system and/or memory please send me a message or a reference on this.
Thanks.
Pete -- Peter Bond Plymouth Electron Microscopy Unit University of Plymouth Drake Circus Plymouth Devon UK PL4 8AA Tel/Fax: 01752 233092 email: pbond-at-plymouth.ac.uk
If you are looking for an inexpensive TEM and/or SEM, The University of Portland has two such units you might be interested in.
1) Zeiss EM9s2, transmission electron microscopy, with additional high voltage tank, replacement for vacuum tubes, fuses, new filaments, lots of film cassettes, and film, original schematics, and operation manuals.
2) ETEC AutoScan, scanning electron microscope, with 60 and 90 degree stages, several reconditioned column liner tubes, new apertures, new YAG scintillator in original container, original schematics and operating manuals, plus, video tapes on operation and use. This instrument was original manufactured for INTEL and has a 50kV power supply.
Both units are operational!
Any interested parties or individuals can contact me off the server.
Ken Tiekotter, Adjunct Professor The University of Portland Department of Biology 5000 N. Willamette Blvd. Portland, OR 97203 USA
EMSL Analytical Inc. (Westmont, NJ) is interested in hiring a Transmission Electron Microscopy Analyst (TEM) for its NJ Corporate Office. EMSL is the world leader in asbestos analysis since 1981 with over 20 laboratory locations worldwide.
Responsibilities include the preparation and analysis of air, bulk, and water samples submitted by clients for asbestos content and other specialty asbestos projects. College Degree in Materials Science or related field and past experience with electron microscopy analysis preferred. . The ideal candidate would be a detail orientated individual able to follow lab protocols and procedures and able to work in a fast paced environment.
Full benefits package with salary commensurate with experience. Interested individuals send resume and salary requirements to Stephen Siegel, CIH (ssiegel-at-emsl.com) or fax to 856-858-4960.
Stephen Siegel Stephen Siegel, CIH Asbestos Lab Manager EMSL Analytical, INC 107 Haddon Avenue Westmont, NJ 08108 Phone:800-220-3675x1209 Fax:609-858-4960 email:ssiegel-at-emsl.com
Your main problem is with the use of SiC for grinding and thinning.
Silicon carbide paper should not be used to prepare any ceramic material, it is not hard enough to cut it efficiently without damaging the substructure. SiC becomes dull quite rapidly, with ceramic a dull abrasive actually cracks the structure and causes pullout and excessive chipping. You are compounding your problem with the weight of the tool being used as well. While the disc grinder is a well designed product and very useful, it is quite heavy for samples of certain thicknesses not to mention the additional pressure you are applying is not helping any.
The solution is to use Diamond Lapping Film and apply some Diamond Extender at the final stages of thinning to reduce the surface tension between the water and the sample. This reduces the sheer stress being applied to the sample and will drastically reduce the possibility of cracking.
Should you have interest, we offer a polishing machine "The MultiPrep System" that allows you to prepare the sample without the possibility of the tool being misaligned or mishandled (tilted on edge) during prep. Only the sample makes contact with the abrasive and the plane of polish remains in tact throughout the polishing/grinding process. A dial indicator (1 micron increments) allows you to preset a known amount of material to be removed and there is no operator intervention until the sample is done. The amount of force applied to the sample is constant regardless of the operator.
If you wish to get more information about the MultiPrep System, you may visit our website: www.alliedhightech.com or contact me in person at 310- 635-2466.
Sincerely,
Gary Liechty Manager, Technical Products Allied High Tech Products, Inc. } ------------------------------------------------------------------------ } 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'm currently having a few problems preparing good cross-section specimens } of ceramic thin films on LaAlO3 or NdGaO3 substrates. } } I can cut them okay, but thinning them often results in cracking as the } thickness goes below 100 microns. I have a Gatan model 623 disk grinder and } currently have access to SiC abrasive papers to grit sizes of 1500 or 2000, } and also have the Gatan diamond polishing disks and specimen lapping kit. } } The problems are as follows: } If we use the 1500 or 2000 grit SiC, we can often get the samples thin } enough, but with poor surface finish which needs to be improved with } dimpling (on both sides), we also risk cracking the sample as SiC papers } often contain imperfections. } } Using the Gatan specimen lapping kit, I find that as the disk grinder is } altered to grind off a further 10 microns, the edge of the sample often } catches on the diamond disk and tears some of the diamond coating off, } leaving a lump on the surface which then risks catching the specimen and } cracking it. } } I have one possible alternative approach which I used in Sweden last year } (with SiAlON ceramics) involving attaching the sample to a glass slide and } polishing it using diamond spray on non-absorbent paper. I may consider } trying this with these new materials. } } Other ideas would be welcome, however, as would suggestions of how to } improve my present method. Please note that I do not have the budget to buy } any expensive new polishing equipment (such as a tripod polisher, for } instance), so the most welcome suggestions would be those that involve } improvements to current techniques, use of different types of polishing } consumables, etc.. } } I hope to hear several suggestions, both from users and from the companies } who producing polishing and lapping equipment. Why not post them with the } list so we can all benefit from the sharing of experience? } } Best wishes } } ===== } Ian MacLaren } Beijing Laboratory of Electron Microscopy } Chinese Academy of Sciences, P.O. Box 2724 } 100080 Beijing } China } General Email: ian.maclaren-at-physics.org } Work (esp. large attachments): maclaren-at-image.blem.ac.cn } } ____________________________________________________________ } Do You Yahoo!? } Get your free -at-yahoo.co.uk address at http://mail.yahoo.co.uk } or your free -at-yahoo.ie address at http://mail.yahoo.ie }
We have JEOL 840 SEM that has develop a leak at the specimen chamber door interface and will not hold a vacuum. We have changed the o-ring around the door, look for nicks, scratches any other damage around the door interface, none found. We have also check the roughing valve (V4, LV3 and the pressure relief valve) for leaks. Pressure holds in the gun area. The large door is only opened when we have to replace or add a new attachment to the chamber and thats about once a year. We have a vacuum specimen exchange port to enter and retrieve samples. We have also check that seal and it also checks out OK. We have had 2 JEOL engineers take a look at the system and at this point no luck finding the cause or solution. You can respond to me off-line at bruce.arey-at-pnl.gov Thanks
hello Kim, Jordi & interested parties, We have a filament ramp circuit on our LaB6 2010 that controls the filament current. It is an accessory offered by Jeol. Jeol has set it to take about 8 minutes to bring the filament to max current. The max. current is still controlled by the (preset) filament current knob on L1. Once the filament is hot, when we change samples or other wise have the filament current off for {5mi. we use the "quick timer" feature of this circuit which brings the filament up in ~90 seconds. The max. filament current is set to run just under saturation (so you can just barely see the X when the beam is converged). The bias control is used to set beam current to 10 uA. The setting changes with KV & filament wear. The initial bias setting is a function of the physical position of the filament relative to the Wehnelt cap. It changes every time the filament or cap are serviced. As the tip wears the difference in bias settings say between 100 & 200 KV increases & both move up. We are a multiple user facility. Our filament sees a of cycles & we always have novice users. Filament life times are 400-1000 hours. This tends to track with the novice user density. As far as the drift Jordi mentioned. This is a non issue here & may be in the filament design. We use the Gimble Phillips 60-06. I do a lot of carbon work & can pretty much nail 3.4A when I get a beam. Yes the images are sharper after we have been running a bit.
Bruce Brinson Rice U.
usual disclaimer... no financial interest in companies mentioned.
Marti, Jordi 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 } -----------------------------------------------------------------------. } } Kim: } } We have a 2010 and our procedure is to bring the filament dial (labeled } off-10) to position #3. Then we go for coffee. After 20 min. or so we raise } it to # 4, after a couple of minutes to #5 etc. until we get to the stop ( } in our case we have it at about 7 1/2). Even at this point we have filament } drift for quite some time, so , we are not ready to take pictures for } another 1/2 hr or so. By the way, we have the bias settings at 7 and 7 which } is high. I would be interested in knowing what your settings are. } } I hope this helps } } Jordi Marti }
A user of our facility is interested in making height measurements from specimens viewed in the SEM. I am aware of the conventional way of doing this: stereo pairs and optical viewer (with stereometer parallax corrections). Is there a more modern (computerized) way of doing this, say with anaglyphs?
Thank you.
John B.
#################################################################### John J. Bozzola, Ph.D., Director Micro-Imaging and Analysis Center 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 Web: http://www.siu.edu/departments/shops/cem.html ####################################################################
In a message dated 6/8/00 12:38:48 PM, bozzola-at-siu.edu writes:
} A user of our facility is interested in making height measurements } from specimens viewed in the SEM. I am aware of the conventional way } of doing this: stereo pairs and optical viewer (with stereometer } parallax corrections). Is there a more modern (computerized) way of } doing this, say with anaglyphs?
There was a thread recently on measurement of stereo pairs by computer-matching of points. Several software packages (one of the is Fovea Pro - http://members.aol.com/FoveaPro) were mentioned as having this capability. Whether or not the two images are put together as an anaglyph is unimportant.
Just a couple of additional comments to Barbara's procedure.
A Zeiss axiom: If the lens is not dirty, then cleaning it will never improve it, it only risks damaging it. Do the least amount possible to return the surface to clean. If the surface is only dusty, & puffing it removes the problem, stop there. I look at the lens being cleaned, using the ocular turned upside down as a loupe, at each step of the process. When it's clean, you're done. Usually only external surfaces are contaminated and need frequent or extensive cleaning. Internal lenses, a dusting often suffices.
In dealing with a dissecting microscope with a zoom magnification capability as opposed to one with click stops or a fixed magnification, be very careful about disturbing the positional settings of the internal lenses. Any changes to these settings will alter magnification for the each of the two eyes and make an image that can't be justified in the brain. This will require a service engineer to remedy. For heavily filmed lenses, use short or broken cotton tipped applicator and get in the best you can. Varying the zoom control will move the lenses up and down and maybe give you enough room to work.
As Barbara wrote, removing loose dust is essential to preserving lens quality. Lenses are coated with various coatings and these are easily damaged. Puffers from camera stores are good. The red bulb ear syringe for babies is excellent and usually readily available. I do use dusters and compressed air although both are frowned upon, by some, as possibly damaging lens coatings. If you use a duster, don't shake or tip it while dusting the lens. This will expel liquid from the can and contaminate the lens worse.
A good lens cleaner that is easy to get is Sparkle Glass Cleaner available from Ace hardware, and grocery stores (No financial interests). Never use Windex. It contains oils that will coat the lens.
As much as possible use the cotton tipped applicators rather than lens tissues. Unless you wear latex or polyethylene gloves, finger oils will get onto the tissue and be transferred to the lens. Another reason to shy away from the tissue is the tendency to scrub the lens. I agree with Barbara, NEVER, NEVER Kimwipes, I've been told from many sources they contain many silica strands, and will scratch the delicate optical coatings. On expensive lenses, make a single pass, using minimal pressure straight across the lens, with the applicator, rolling the stick between your fingers to present a fresh cotton surface to the lens and picking up and getting the dust away from the lens. Alternate between wet (either with lens cleaner or condensed breath (open mouth, no spit please) and dry cotton. (Yes, it is often necessary to use a swirling motion on oculars because of the amount of oil contamination from the eye lashes.)
It is best not to try to clean filters, there are a few recommended procedures but all potentially damage the very precise and thin coatings of the filter compromising its performance. (If anyone disagrees or has a favorite procedure for filter cleaning, contact me off list, I'd like to hear about it.)
Use of solvents or disassembling objectives or multiple lens stacks is best left to the service engineer. Lenses use a variety of air and cement interfaces to achieve resolution and aberration correction. Altering these by dissolving the cement will degrade the lens. Getting the small lenses back in exactly the same position is also very, very difficult and not for the faint hearted.
There is a microscope repair workshop in the initial planning stages for the Long Beach MSA meeting next year. It will target K-12 school teachers, but will hopefully address many levels. If members of the list will be at the meeting and are interested in attending this workshop to: 1. learn basic repairs for their microscopes. 2. get ideas for their outreach program. or 3. assist with putting on the workshop. Please e-mail me telling me of your interest or asking for more information.
If you have any questions feel free to contact me off line.
Jim is absolutely right about LR white. But instead of increasing the GA I will probably increase paraformaldehyde conc. to 4%.
Good luck,
Soumitra
} } Hi, } } } } I'm trying to do immunogold studies on a membrane protein in a preparation } } of plant vesicles (from minus 80oC storage). I did an Epon embedding with } } standard procedure (glut and osmium) to check the ultrastructure and got } } decent-looking vesicles, but side by side an LR White prep (0.5% glut, 2% } } paraformaldehyde, without osmium) looks like a disaster, with granular } } material, vague membranish-like structures and whorls of membranes. Does } } anyone have any tips for LR White preps of vesicles, or any other ideas } } for embedding (resins, fixing, dehydration, embedding) to favour the } } immunogold process. } } } } Thanks, } } } } Mark } } } } } } } } } } } } } } ******************************************** } } Mark West, } } Unidad de Microscopia Electronica, } } (Electron Microscopy Unit) } } Instituto de Fisiologia Celular, } } Universidad Nacional Autonoma de Mexico, } } 04510 Mexico D.F. } } } } tel (unidad/lab) *(525) 622 5610* } } (casa/home) (525) 619 3020 } } Fax (525) 616 2282 } } ******************************************** } }
***************************************************************** Soumitra Ghoshroy Ph.D. Electron Microscopy Lab Box 3EML New Mexico State University Las Cruces, NM 88003 Tel: 505-646-3600 Fax: 505-646-5665 e-mail: ghoshroy-at-nmsu.edu http://confocal.nmsu.edu/eml
} ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. text deleted } } It is best not to try to clean filters, there are a few recommended } procedures but all potentially damage the very precise and thin coatings of } the filter compromising its performance. (If anyone disagrees or has a } favorite procedure for filter cleaning, contact me off list, I'd like to } hear about it.) } Please respond on-line! I'm sure that I'm not the only other person who would be interested in this!
Try treat the glut- & Os-fixed sections you have with sodium metaperiodate before doing immunolabeling. It may work.
Reference: M. Bendayan 1989. Protein A-gold and Protein G-gold postembedding immunoelectron microscopy. In Colloidal Gold: Principles, Methods and Applications Vol.1 Academic Press.
Ann Fook Yang EM Unit, Eastern Cereal and Oilseed Research Centre, Rm 2091, K.W. Neatby Bldg., Central Experimental Farm, Ottawa, Ontario, Canada K1A 0C6
Phone: 613-759-1638 Fax; 613-759-1701
} } } Mark West {mwest-at-ifcsun1.ifisiol.unam.mx} 06/07 11:50 AM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi,
I'm trying to do immunogold studies on a membrane protein in a preparation of plant vesicles (from minus 80ĽC storage). I did an Epon embedding with standard procedure (glut and osmium) to check the ultrastructure and got decent-looking vesicles, but side by side an LR White prep (0.5% glut, 2% paraformaldehyde, without osmium) looks like a disaster, with granular material, vague membranish-like structures and whorls of membranes. Does anyone have any tips for LR White preps of vesicles, or any other ideas for embedding (resins, fixing, dehydration, embedding) to favour the immunogold process.
Thanks,
Mark
******************************************** Mark West, Unidad de Microscopia Electronica, (Electron Microscopy Unit) Instituto de Fisiologia Celular, Universidad Nacional Autonoma de Mexico, 04510 Mexico D.F.
Sorry, I don't have any useful suggestions, and for that reason, I'd rather see responses posted to the list, if possible, as this gives me a wonderful and otherwise unavailable opportunity to learn from the experience of others.
cheers
rtch
} ------------------------------------------------------------------------ } 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 have JEOL 840 SEM that has develop a leak at the specimen chamber door } interface and will not hold a vacuum. We have changed the o-ring around the } door, look for nicks, scratches any other damage around the door interface, none } found. We have also check the roughing valve (V4, LV3 and the pressure relief } valve) for leaks. Pressure holds in the gun area. The large door is only opened } when we have to replace or add a new attachment to the chamber and thats about } once a year. We have a vacuum specimen exchange port to enter and retrieve } samples. We have also check that seal and it also checks out OK. } We have had 2 JEOL engineers take a look at the system and at this point no luck } finding the cause or solution. You can respond to me off-line at } bruce.arey-at-pnl.gov } Thanks } } Bruce W. Arey } } PNNL-Battelle } Associate Scientist } Microstructural Characterization } bruce.arey-at-pnl.gov } 509-376-3363 } fax 509-376-6308 } } } } 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
Mark, you didn't say how you post-stained after you immunolabeled. This step can make all the difference in the world. For LRW, I prefer 3-4 secs in saturated UA in 50% etoh, wash, followed by about 15secs in lead citrate. I find a major difference (inferior) when I use aqueous UA. Also, going too long in any of the solutions will give poor results.
} ===== Original Message From Mark West {mwest-at-ifcsun1.ifisiol.unam.mx} ===== } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hank Adams Manager Integrated Microscopy Core Molecular and Cellular Biology Baylor College of Medicine Houston, Tx 77030
John - I routinely make height measurements of 5 - 10 um interplanetary dust particles using the objective focus knob on our JEOL 1200EX in STEM mode. The procedure is simply to focus on the top of the particle and note the number of steps (or count the number of clicks of the knob) required when refocussing on the substrate next to the particle. By calibrating the objective focus step size with a known standard, you can routinely measure particle heights. In principle, a similar procedure should be possible on an SEM, depending on how the objective focus is configured. I don't think you can expect high accuracy with this method, however, so it may not be applicable to your needs.
Dave Joswiak Dept. of Astronomy University of Washington Seattle, WA 98195 joswiak-at-astro.washington.edu
On Thu, 8 Jun 2000, John J. Bozzola 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 } -----------------------------------------------------------------------. } } } A user of our facility is interested in making height measurements } from specimens viewed in the SEM. I am aware of the conventional way } of doing this: stereo pairs and optical viewer (with stereometer } parallax corrections). Is there a more modern (computerized) way of } doing this, say with anaglyphs? } } Thank you. } } John B. } } } #################################################################### } John J. Bozzola, Ph.D., Director } Micro-Imaging and Analysis Center } 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 } Web: http://www.siu.edu/departments/shops/cem.html } #################################################################### } }
Tamara et al : One superior method for cleaning sensitive optical components such as filters and front surfaced telescope mirrors is as follows: 1) gently puff of extraneous dust. 2) gently float the surface with deionized water. 3) if necessary to remove dirt film use a q-tip soaked in a non-ionic surfactant like a 0.01% solution of triton-x 100 (IMPORTANT DO NOT RUB ON THE SURFACE WITH FORCE BUT SIMPLY LET THE WEIGHT OF THE WET Q-TIP(S) BE THE ONLY "DOWNWARD PRESSURE" ON THE SURFACE OF THE GLASS. Such gently techniques such as these should help mitigate damage.
Tamara et al : (excuse mispelling in first copy sent) One superior method for cleaning sensitive optical components such as filters and front surfaced telescope mirrors is as follows: 1) gently puff off extraneous dust. 2) gently float the surface with deionized water. 3) if necessary to remove dirt film use a q-tip soaked in a non-ionic surfactant like a 0.01% solution of triton-x 100 (IMPORTANT DO NOT RUB ON THE SURFACE WITH FORCE BUT SIMPLY LET THE WEIGHT OF THE WET Q-TIP(S) BE THE ONLY "DOWNWARD PRESSURE" ON THE SURFACE OF THE GLASS. Such gently techniques such as these should help mitigate damage.
} I remember a message some time ago, I think, suggesting a way to flat } embed in silicon molds using LRWhite resin. How the top was sealed } escapes me. Could anyone with a better filing system and/or memory } please send me a message or a reference on this.
Peter Bond {P.Bond-at-plymouth.ac.uk}
} Pete -
If you have a silicon mold that isn't permeable to LR White, I'd like to know who makes it! There is a teflon flat mold available from Ted Pella that works well with LR White; it's their own product. It's sealed with Thermanox coverslips, so you might try them with your mold.
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
Peter Bond wrote: ============================================================== I remember a message some time ago, I think, suggesting a way to flat embed in silicon molds using LRWhite resin. How the top was sealed escapes me. Could anyone with a better filing system and/or memory please send me a message or a reference on this. ============================================================== This might have been an old posting of mine. Since UV transparency is usually required, we are talking about clear UV transparent silicone embedding molds and we recommend a slight "overfilling" of the cavities. Then when all cavities are filled, take another mold, just like the first one, and place it on top, bottom side down, cavity side up. The capillary action will result in a sealing out of any oxygen.
When the UV curing is complete, the top mold can be easily separated and since the cavity side is still unused, no wear and tear has been put on it in terms of taking away any of its lifetime.
Information about these transparent-to-UV embedding molds, and their use, can be found on the SPI website given below.
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.spi.cc ######################## ============================================
Dear colleagues, We have a problem with spectra recording in our EDAX DX4 system mounted on Philips CM12/STEM. There is a hole in the spectrum between 0.5keV and 3.5keV. With service engineer, we have borrowed all the boards in electronics and exchange our boards with the borrowed ones. The hole in the spectrum remains. We have also dismounted detector from the EM column and the Be window was checked for contamination. The window was clean and intact. Please, can anybody give us any hints how to solve our problem? Many thanks in advance for any comment. Oldrich Benada
Our system specification: Philips CM12/STEM EDAX DX4 system with 184 preamp. based on win3.11
+-----------------------------------+ Oldrich Benada Acad. Sci. CR Institute of Microbiology Laboratory of electron microscopy Videnska 1083 CZ - 142 20 Prague 4 - Krc Czech Republic +------------------------------------+ Phone: +420-2-4752399 Fax: +420-2-4715743 WEB:http://www.biomed.cas.cz/mbu/lem113/lem.htm
Balzers Union renamed to BAL-TEC AG in 1992. But the product line was maintained and new developments are carried out. For additional information have a look at our website www.bal-tec.com. There are several coating systems and freeze fracture systems of this older types in use.
G'day Folks, I acquired some carbonate standards a while ago from the Smithsonian Insitute. Namely, Calcite, Siderite, Dolomite and Strontianite. Interestingly, when admiring them with our ED system, there appeared an anomalous peak around about where B is supposed to be. This is not a detector artifact as it is specific only to these materials. I just wondered if anyone else had noticed the same thing, or whether anyone has any clues on why this should be occuring. The programme suggests that there is about 50 wt % B in these things which seems unbelievable considering the compositions supplied by Washington. Ideas? Cheers, Malc.
-- Dr MP Roberts Phone: [+27](0)46 603 8313 Dept of Geology Fax: [+27](0)46 622 9715 Rhodes University Cell: 083 4060 262 (usually off) 6140 Grahamstown e-mail: malc-at-rock.ru.ac.za SOUTH AFRICA
With regard to the vacuum problems encountered by Bruce , without being familiar with the instrument we encountered a ' similar ' problem and were misled into thinking a chamber door leak existed by an over zealous leak detection unit . The problem on our sem actually turned out to be a hairline crack in a metal bellows attached to a valve that appeared OK . The vaccum would not exceed a certain level in pumpdown in its final stages .
I believe these bellows were often a source of problems on older instruments .
M.HARRIS harrism-at-esm-semi.co.uk ESM LTD South Wales , U.K .
Hi All, I'm posting this for a friend who is not on-line.
Does anyone out there have a used RMC MT-7000 for sale? My friend just moved to a lab that had an MT-5000 which she was told was operational. A major exaggeration. She is an RMC loyalist, and desparately wants another, but has a limited budget.
Please contact: Linda Burg Friedman at Columbia P & S at (212)305-9047
Thanks, Lee
Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Confocal Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
The specimen chamber is a vast area with many potential leak sources.
The door is the most likely if it is being opened on a regular basis, however do not forget that the stage drives pass through the door and they are being used all of the time.
If you are sure the door "O" ring is OK check the stage drive feed throughs as they may be the source of your problem?
Steve Chapman Senior Consultant Protrain For consultancy and professional training in EM world wide Tel 44+ 1280 814774 Fax 814007 www.emcourses.com
Rick, YOu don't mention make or modle of TEM, but i can tell you that my service contract (2 preventive maintenance call, unlimited service calls, parts, labor) costs around $15,500/year. You will need a water supply for cooling and electrical work to bring in a dedicated line. You will need a dakrroom with running water and a temp. control valve to process the film. If you wish to make photographic prints of your negatives, you will also need a point source enlarger (Durst was the best, but they are hared to find, Omega used to make one, too) and either pans (slow and painful) or a rapid processor. Budgeting for supplies is a tough call...it depends on your usage. If you find someone with an EM background to run it, its should't take them long to learn the individual instrument. Training someone from ground zero could take months to get dthe person on his/her way. Ancillary euqipment: it depends on what you will be doing, biological or materials, embedding/sectioning or particulates, negataive staining or metal shadowing.
A really good source to check is Audrey Glauert's Practical Methods in Electron Microscopy Vol. 4: Design of the Electron Microscope Laboratory, by Ronald H. Alderson, North-Holland publishers, 1975.
If you wish to contact me off-line, I can go over my lab's operating budget with you. I run a basic EM Core Facility here at (what used to be called) Cornell Medical College.
Lee
Leona Cohen-Gould, M.S. Sr. Staff Associate Director, Electron Microscopy Core Facility Manager, Confocal Microscopy Core Facility Joan & Sanford I. Weill Medical College of Cornell University voice (212)746-6146 fax (212)746-8175
JEOL has been very helpful and they are somewhat limited on what they can do to this instrument. The 840 has radioactive particles inside the chamber so we have to be careful each time we open the system up plus there hands on is limited to just oversight. I am in the process of trying to find a leak detector on site that can be used on radioactive system. But we also cannot pump down the chamber enough to use a leak detector. So we are in the process of making a plate that will fit over the door. We have taken off all our accessories off the chamber (EDS, OIM, BSE) and have pressurized the system and have found some evidence of a leak on the left hand side on the door, we have changed the o-ring and still the seal leaks on this side of the door. We have polished the door seal to make sure there is no major scratches or marks. We are pretty confident that specimen exchange port is OK we can pump this down and we see no signs of a leak. Why are we focusing our attention on the large o-ring and chamber door. We can clean the o-ring with alcohol (methanol or ethanol) and we can rough pump the chamber out and put the chamber on the high vac system but after a period of time the alcohol dries out and the system shuts down too a poor vacuum (overnight). Then we try to rough pump the chamber and we cannot go more than 20-30 mamps difference on the pirani gauge. We have done this several times with the same results. We are going to try to pressurize the system with He and try He sniffer to help us maybe pin point the leak. Thanks to all who have responded with suggestion on finding the leak and if we are successful in finding the leak I will post the finding on the server. Again JEOL has been very responsive in trying to find the leak and they are somewhat limited on this instrument do to its environment. Any other suggestion are welcomed.
---------- From: "Dopeyee-at-aol.com"-at-sparc5.microscopy.com Sent: Thursday, June 8, 2000 4:28 PM To: Microscopy-at-sparc5.microscopy.com Subject: LEAK IN JEOL
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GET A HOLD OF A LEAK DETECTOR AND FIND THE TRUE AREA THAT IS LEAKING
I DO BELEIVE THE MANUFACTURER SHOULD BE OF MORE HELP
I recall many years ago we had a similar vacuum leak on our 840A that was related to the specimen exchange port mechanism. The moving parts in the sliding door mechanism had become slightly miss-aligned. We ended up disassembling that mechanism, installing new o-rings and lubricating with Apiezon L. Problem gone. Hope your leak is that easy! Good Luck Brad Huggins BP Amoco, Naperville
} ---------- } From: Arey, Bruce W[SMTP:bruce.arey-at-pnl.gov] } Sent: Thursday, June 08, 2000 9:24 AM } To: 'Microscopy-at-msa.microscopy.com' } Subject: JEOL 840 SEM Vacuum Problem } } ------------------------------------------------------------------------ } 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 have JEOL 840 SEM that has develop a leak at the specimen chamber door } interface and will not hold a vacuum. We have changed the o-ring around } the } door, look for nicks, scratches any other damage around the door } interface, none } found. We have also check the roughing valve (V4, LV3 and the pressure } relief } valve) for leaks. Pressure holds in the gun area. The large door is only } opened } when we have to replace or add a new attachment to the chamber and thats } about } once a year. We have a vacuum specimen exchange port to enter and retrieve } samples. We have also check that seal and it also checks out OK. } We have had 2 JEOL engineers take a look at the system and at this point } no luck } finding the cause or solution. You can respond to me off-line at } bruce.arey-at-pnl.gov } Thanks } } Bruce W. Arey } } PNNL-Battelle } Associate Scientist } Microstructural Characterization } bruce.arey-at-pnl.gov } 509-376-3363 } fax 509-376-6308 } } }
I have always understood that deionized water would etch metal, that being the reason for PVC pipe being used to deliver it. If this is true, wouldn't that damage the first surface mirrors?
David, We have a B&L StereoZoom (0.7X - 3X) microscope which we intend to donate to the local school system (Computer VoTech Dept) for use in examining the circuit patterns on silicon wafers that I have donated to them.
My problem is that the microscope is not functioning properly. The two optical paths are not synchronized during zooming and therefore the image tends to rotate or it falls out of focus. We have tried to repair it in-house but we were unsuccessful. Knowing that that we would create additional problems, we did not disturb the lens or prism sub-assembly.
Is there a set of maintenance instructions or a tech manual available to help us align this scope?
Thanks Mike Urbanik Commercial Crystal Labs Naples, FL www.crystalguru.com
you may want to try the method below if your samples show very large structures. One of the nice features of an SEM is it's large depth of focus, unfortunately in many cases this prevents you from using the technique mentioned below. You can use a stereo technique to calculate a surface profile (see for example the stereo module on our web site or other stereo applications). Typical results from stereo images have a height resolution of about 1/10 the of the lateral resolution (in other words: if your lateral resolution is 1 micron between pixels, the height resolution will be on the order of 10 microns). This can be improved by sub-pixel interpolation but gives you an order of magnitude for the resolution.
Michael
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-----Original Message----- } From: David Joswiak [mailto:joswiak-at-orca.astro.washington.edu] Sent: Thursday, June 08, 2000 4:44 PM To: John J. Bozzola Cc: Microscopy-at-sparc5.microscopy.com
John - I routinely make height measurements of 5 - 10 um interplanetary dust particles using the objective focus knob on our JEOL 1200EX in STEM mode. The procedure is simply to focus on the top of the particle and note the number of steps (or count the number of clicks of the knob) required when refocussing on the substrate next to the particle. By calibrating the objective focus step size with a known standard, you can routinely measure particle heights. In principle, a similar procedure should be possible on an SEM, depending on how the objective focus is configured. I don't think you can expect high accuracy with this method, however, so it may not be applicable to your needs.
Dave Joswiak Dept. of Astronomy University of Washington Seattle, WA 98195 joswiak-at-astro.washington.edu
On Thu, 8 Jun 2000, John J. Bozzola 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 } -----------------------------------------------------------------------. } } } A user of our facility is interested in making height measurements } from specimens viewed in the SEM. I am aware of the conventional way } of doing this: stereo pairs and optical viewer (with stereometer } parallax corrections). Is there a more modern (computerized) way of } doing this, say with anaglyphs? } } Thank you. } } John B. } } } #################################################################### } John J. Bozzola, Ph.D., Director } Micro-Imaging and Analysis Center } 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 } Web: http://www.siu.edu/departments/shops/cem.html } #################################################################### } }
This may sound like "bucket science" but we've used the disposable aluminum weighing dishes to flat embed material. If you fill the bottom tray about 1/2 full and set another tray inside it, press gently until a little LR white oozes up the sides, what remains in the middle will be protected from the air and should polymerize nicely. (Maybe we've just been lucky!) good luck!
Tracey Pepper Supervisor Bessey Microscopy Facility Iowa State University ph: 515-294-3872 fax: 515.294.1337
Some Chlorine-L lines, as well as Sr lines have nearly identical energy/wavelength as B Ka. I don't remember the chemistry of these standards offhand, but I know some of them were Sr-bearing and some may also have Cl. These are likely the peaks you are seeing at the low end.
Jim -- ^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^^ James J. McGee (email: jmcgee-at-sc.edu) Department of Geological Sciences University of South Carolina Columbia, SC 29208
Tel: 803-777-6300 Fax: 803-777-6610
Dr Malcolm Roberts 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 } -----------------------------------------------------------------------. } } G'day Folks, } I acquired some carbonate standards a while ago from the Smithsonian } Insitute. Namely, Calcite, Siderite, Dolomite and Strontianite. } Interestingly, when admiring them with our ED system, there appeared an } anomalous peak around about where B is supposed to be. This is not a } detector artifact as it is specific only to these materials. I just } wondered if anyone else had noticed the same thing, or whether anyone } has any clues on why this should be occuring. The programme suggests } that there is about 50 wt % B in these things which seems unbelievable } considering the compositions supplied by Washington. Ideas? } Cheers, } Malc. } } -- } Dr MP Roberts Phone: [+27](0)46 603 8313 } Dept of Geology Fax: [+27](0)46 622 9715 } Rhodes University Cell: 083 4060 262 (usually off) } 6140 Grahamstown e-mail: malc-at-rock.ru.ac.za } SOUTH AFRICA
If the instrument has been physically moved recently, this can sometimes
hasten a crack in these stainless steel "flex" vacuum lines. I have found these to develop mostly at the end of the tube where it has been flared for the fitting connector. Check the roughing lines first.
Happy hunting...
Bob Roberts EM Lab Services, Inc 2409 S. rural Rd Suite C Tempe, Arizona 85282 480.967.3946
} Oldrich Need a little more information. Do you actually have spectral information in the spectrum at energies { 0.5, or is there possibly, only noise in this low energy range? If the signal present in your spectra at these very lowest energies is just noise-like signal. Then it is possible that you have a severe alignment problem with the EDS detector/specimen geometry. A combination of high noise (due to vibration or other sources) and poor line of sight with the detector window (resulting in detection of only the higher energy x-rays ) could give you this "hole in the EDS spectrum" effect. A miss-aligned detector, and/or a detector making contact with the internal parts of the microscope might create this situation. Does the information in the low energy region correlate with the specimen composition? Does the high energy signal correlate with the specimen composition?
} ---------- } From: Oldrich Benada[SMTP:benada-at-biomed.cas.cz] } Sent: Friday, June 09, 2000 2:19 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: A hole in the EDS spectrum } } ------------------------------------------------------------------------ } 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 colleagues, } We have a problem with spectra recording in our EDAX DX4 system } mounted on Philips CM12/STEM. There is a hole in the spectrum between } 0.5keV and 3.5keV. } With service engineer, we have borrowed all the boards in electronics } and exchange our boards with the borrowed ones. The hole in the } spectrum remains. We have also dismounted detector from the EM column } and the Be window was checked for contamination. The window was clean } and intact. } Please, can anybody give us any hints how to solve our problem? Many } thanks in advance for any comment. } } Oldrich Benada } } Our system specification: } Philips CM12/STEM } EDAX DX4 system with 184 preamp. based on win3.11 } } } } } } +-----------------------------------+ } Oldrich Benada } Acad. Sci. CR } Institute of Microbiology } Laboratory of electron microscopy } Videnska 1083 } CZ - 142 20 Prague 4 - Krc } Czech Republic } +------------------------------------+ } Phone: +420-2-4752399 } Fax: +420-2-4715743 } WEB:http://www.biomed.cas.cz/mbu/lem113/lem.htm }
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I have found the use of clear silcone sealant (as used for windows etc) quite effective to temporally fix or help to isolate large and medium vacuum leaks. In many instance it can be applied externally over various fittings. Screw holes are best covered first with a little tape, to facilitate the later the removal of the dry silicone. Silicone outgases a fair bit for some hours, so only major leaks can be determined immediately after applying the silicone. The method seems crude but is effective to eliminate numerous fittings as the source of a leak. I once operated a TEM for several months with a split stainless bellow, patched with a smear of silicone sealant. I don't suggest the use of that sealant on a permanent basis or in ion gutter pumped parts of a column. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Saturday, June 10, 2000 12:05 AM, Arey, Bruce W [SMTP:bruce.arey-at-pnl.gov] wrote: } } } JEOL has been very helpful and they are somewhat limited on what they can do } to } this instrument. The 840 has radioactive particles inside the chamber so we } have } to be careful each time we open the system up plus there hands on is limited } to } just oversight. I am in the process of trying to find a leak detector on site } that can be used on radioactive system. But we also cannot pump down the } chamber } enough to use a leak detector. So we are in the process of making a plate } that } will fit over the door. We have taken off all our accessories off the chamber } (EDS, OIM, BSE) and have pressurized the system and have found some evidence } of } a leak on the left hand side on the door, we have changed the o-ring and } still } the seal leaks on this side of the door. We have polished the door seal to } make } sure there is no major scratches or marks. We are pretty confident that } specimen } exchange port is OK we can pump this down and we see no signs of a leak. Why } are } we focusing our attention on the large o-ring and chamber door. We can clean } the o-ring with alcohol (methanol or ethanol) and we can rough pump the } chamber } out and put the chamber on the high vac system but after a period of time the } alcohol dries out and the system shuts down too a poor vacuum (overnight). } Then } we try to rough pump the chamber and we cannot go more than 20-30 mamps } difference on the pirani gauge. We have done this several times with the same } results. We are going to try to pressurize the system with He and try He } sniffer } to help us maybe pin point the leak. Thanks to all who have responded with } suggestion on finding the leak and if we are successful in finding the leak I } will post the finding on the server. Again JEOL has been very responsive in } trying to find the leak and they are somewhat limited on this instrument do } to } its environment. Any other suggestion are welcomed. } } } Bruce W. Arey } } PNNL-Battelle } Associate Scientist } Microstructural Characterization } bruce.arey-at-pnl.gov } 509-376-3363 } fax 509-376-6308 } } } ---------- } From: "Dopeyee-at-aol.com"-at-sparc5.microscopy.com } Sent: Thursday, June 8, 2000 4:28 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: LEAK IN JEOL } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } GET A HOLD OF A LEAK DETECTOR AND FIND THE TRUE AREA THAT IS LEAKING } } I DO BELEIVE THE MANUFACTURER SHOULD BE OF MORE HELP } } THEY HAVE TO FIX IT!!!! }
Hmmm, and "I didn't even know that", but believed that deionised water had metal ions removed from it and so in that respect its purer than 2x glass distilled water. Then I was taught and believed! that metal pipes would re-introduce metal ion back into the water! Being deionised the water has no buffering capacity and therefore is neither acid nor alkaline, they told me and I believed. Education is expensive; must ask for my money back. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Saturday, June 10, 2000 12:24 AM, "milesd-at-us.ibm.com"-at-sparc5.microscopy.com [SMTP:"milesd-at-us.ibm.com"-at-sparc5.microscopy.com] wrote: } } } I have always understood that deionized water would etch metal, } that being the reason for PVC pipe being used to deliver it. If this } is true, wouldn't that damage the first surface mirrors? } } Darrell }
Ahhh, but I have now been educated! Re-contamination of the painstakingly purified water is the concern, and there is no threat to the durability of the pipes. I had been mislead.
Hi Vacuum leaks, what a pleasure! Our tried and tested methods include using Petroleum Ether or Ethanol and then Bostick Prestic or Blue tac as it is known in Australia. Spraying alcohol around the suspected areas should show up the leak. If you ant to try and stop a leak use the Prestic. Its that stuff you buy at the stationary shop that is used to stick posters and pictures to a wall. That is pliable, removable and really handy at sealing off a few suspect areas. We are currently working on a JEOL840 here with a leak. After many hours we found that it was the gauge head that was leaking.
Good luck Luc Harmsen Anaspec, South Africa Technical support on microscopy. Tel + 27 (0) 11 476 3455 www.anaspec.co.za
ICEM 15 will be in Durban, South Africa, 2002. www.icem15.com
-----Original Message----- } From: Arey, Bruce W [mailto:bruce.arey-at-pnl.gov] Sent: Friday, June 09, 2000 4:05 PM To: 'microscopy-at-msa.microscopy.com'
JEOL has been very helpful and they are somewhat limited on what they can do to this instrument. The 840 has radioactive particles inside the chamber so we have to be careful each time we open the system up plus there hands on is limited to just oversight. I am in the process of trying to find a leak detector on site that can be used on radioactive system. But we also cannot pump down the chamber enough to use a leak detector. So we are in the process of making a plate that will fit over the door. We have taken off all our accessories off the chamber (EDS, OIM, BSE) and have pressurized the system and have found some evidence of a leak on the left hand side on the door, we have changed the o-ring and still the seal leaks on this side of the door. We have polished the door seal to make sure there is no major scratches or marks. We are pretty confident that specimen exchange port is OK we can pump this down and we see no signs of a leak. Why are we focusing our attention on the large o-ring and chamber door. We can clean the o-ring with alcohol (methanol or ethanol) and we can rough pump the chamber out and put the chamber on the high vac system but after a period of time the alcohol dries out and the system shuts down too a poor vacuum (overnight). Then we try to rough pump the chamber and we cannot go more than 20-30 mamps difference on the pirani gauge. We have done this several times with the same results. We are going to try to pressurize the system with He and try He sniffer to help us maybe pin point the leak. Thanks to all who have responded with suggestion on finding the leak and if we are successful in finding the leak I will post the finding on the server. Again JEOL has been very responsive in trying to find the leak and they are somewhat limited on this instrument do to its environment. Any other suggestion are welcomed.
---------- From: "Dopeyee-at-aol.com"-at-sparc5.microscopy.com Sent: Thursday, June 8, 2000 4:28 PM To: Microscopy-at-sparc5.microscopy.com Subject: LEAK IN JEOL
------------------------------------------------------------------------ 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 -----------------------------------------------------------------------.
GET A HOLD OF A LEAK DETECTOR AND FIND THE TRUE AREA THAT IS LEAKING
I DO BELEIVE THE MANUFACTURER SHOULD BE OF MORE HELP
We are consolidating two electron microscopy labs in the Boston area and have some equipment which someone may want. 1) JEOL JEM 100CX electron microscope which has always been under service contract and is still being used. It is about 20 years old and we would like to see it in a new home rather than trashing it. You would need to have it moved. 2)JEOL JEE 4C vacuum evaporator for Carbon. Still under vacuum and yours for the taking. 3)Durst Laborator 138S floor model enlarger with many condensers and an Agfa Rapidoprint DD6400 processor, both in very good shape and for sale. Contact me directly with any questions.
Norman Michaud Director, Morphology Mass Eye and Ear Infirmary Ophthalmology-5th flr. 243 Charles St, Boston, MA 02114 norman_michaud-at-MEEI.Harvard.edu Tel:617-573-3316; Fax:617-573-4290
Bruce W. Arey referred in his request to find a vacuum leak in his JSM840 to 'radioactive particles' inside the chamber of his SEM. I wonder if he could be more specific, because I feel if we start calling a SEM as a 'radioactive system' many safety officers will have a field day. I have been working with various SEMs since 1968 and never felt that I might be bombarded by radioactive particles, even when I opened the chamber (e.g. ETEC) of the SEM.
Hans Brinkies Senior Lecturer Swinburne, University of Technology School of Engineering and Science P.O.Box 218 - Hawthorn - Vic -3122 - Australia Phone: +61 3 9214 8657 Fax: +61 3 9214 8264 Email: Hbrinkies-at-swin.edu.au
I have a student who has been trying to jet polish tungsten for TEM. She has tried various concentrations of sodium hydroxide in water, as well as 40g trisodium phosphate/250ml water, and 55.8g magnesium perchlorate/250ml methanol, at a range of voltages. We have a Fischione jet-polishing unit. So far, none of the samples has been close to good. Can anyone help us out here, with past experience or general suggestions?
Many thanks in advance,
Gill
Dr Gillian M. Bond Department of Materials & Metallurgical Engineering New Mexico Tech Socorro, NM 87801
Certainly the best reference you can have for any jet polishing inquiry is Bernie Kestel at Argonne National Laboratory. I will forward this to him to see if he can add anything else. In digging through my extensive "Bernie Archives" I did find a paper titled "A Jet Polishing Solution for Silicon Germanium, Tantalum, Niobium and Tungsten-Rhenium" Ultramicrscopy 9 (1982) 379-384.
He was able to get a good polish under the following conditions: Temperature: -50 C Jet Height: 4.5mm (Single vertical jet system) Pump setting: 6 Volts: 40 Current: 20mA
This was done using his BK-1 solution. BK-1 is prepared by mixing 500ml methanol, 100ml butyl cellosolve, 90ml H2SO4 and 30ml HF.
He also has another paper MRS Volume 199 "Improved Methods and Novel Techniques for Jet Electropolishing of TEM Foils" which lists a method for electropolishing a 0.010" tungsten wire.
He was able to get a good polish under the following conditions: Temperature: -50 C Pump setting: 6 Volts: 120V
Using 6% HF, 12% sulphuric acid, 68% methanol, and 14% butyl cellosolve.
Of course these were done with a South Bay Vertical Jet system so you will need to adjust the parameters for your system. Please get the reference papers or contact me and I will send them to you. I have a long list of Bernie's papers I could send you along with many other references on TEM sample preparation that may be of interest. If you'd like to see more, please contact me.
DISCLAIMER: South Bay Technology produces the Model 550 D Single Vertical Jet Electropolisher as described above and, therefore, has a vested interest in promoting its use.
David Henriks Vice President TEL: 800-728-2233 (toll free in the USA) South Bay Technology, Inc. +1-949-492-2600 1120 Via Callejon FAX: +1-949-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy.
Message text written by Gillian Bond } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I have a student who has been trying to jet polish tungsten for TEM. She has tried various concentrations of sodium hydroxide in water, as well as 40g trisodium phosphate/250ml water, and 55.8g magnesium perchlorate/250ml methanol, at a range of voltages. We have a Fischione jet-polishing unit. So far, none of the samples has been close to good. Can anyone help us out here, with past experience or general suggestions?
Many thanks in advance,
Gill
Dr Gillian M. Bond Department of Materials & Metallurgical Engineering New Mexico Tech Socorro, NM 87801 {
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Business continuity and disaster recovery planning has now been made easier than ever with Version 7.3 of the Disaster Recovery System (DRS) product. DRS provides a plan for inaccessibility or inoperability (a disaster situation).
DRS is an industry standard software product, used by thousands worldwide. DRS users are in large and small companies across a wide variety of industries.
DRS conforms to federal regulations and meets insurance, auditing and legal requirements. DRS runs under Windows, with stand-alone and network versions available. DRS provides the most complete, easy-to-use product available today.
To prove its value, a free trial is available. For more information, visit our web site at www.drsbytamp.com {http://www.drsbytamp.com} . Dealer and distributor inquiries are welcomed
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Hi, one of our TEM's , a Zeiss EM 109, is equipped with an ion getter hi-vac pump. We are now planning to install a turbopump including all the piping and valves that are necessary. The reason for this is that the high vacuum is not good enough ( only in low E-6 area ) and the ion pump has to work too hard and the life-time of the electrodes becomes very short. Also the housing is clogged with trapped gas molecules and has to be regenerated far too often. Technically and electronically this exchange is fairly easily done, but my question is if anyone has done it and if so, what's the experience? Yours sincerely
Per Hšrstedt Department of Medical Biosciences Pathology Unit for Electron Microscopy University of UmeŚ S-90187 UmeŚ Sweden
Dear Friends, For a long time we used Fab fragments-HRP from BioSys (France) for the immuno EM labelling of proteins with subsequent preembedding. These fragments were the best. However, recently the company cancelled its activity and does not send any more these products. Would you be so kind to tell me what has happened (if you know) and what fragments have the same quality?
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I changed my former SEM (Etec w/D.P)to a a Leybold mag-lev turbo a number of years ago. The ball bearing pumps (of the day) caused too much vibration. The results were excellent! The Etec plumbing allowed the turbo to run continuously during vent cycles which was of great benefit.
Keep in mind that for any gas, turbos have a fixed compression ratio. Among other things, the foreline pressure will have a direct influence on the ultimate vacuum.
The ion pumos are better than turbos for the cleanest, highest vacuum, but you have observed one weakness. They do not excel at pumping large volumes of garbage-loaded gasses.
Good Luck,
Woody White McDermott Technology
Hi, one of our TEM's , a Zeiss EM 109, is equipped with an ion getter hi-vac pump. We are now planning to install a turbopump including all the piping SNIP
I believe The SEM to which Bruce refered was/is used to examine radioactive materials. The "zoomies" are not from the SEM itself, but contamination from his specimens. My (former) Etec was in a similar condition. Over the years, my work mix resulted in a stage/chamber activitly level in the thousands of cpm generated by radioactive products of nuclear fission. ...Kept the really loose stuff at a minimum, but certainly had to exercise the appropriate precautions when working on the system.
Woody White McDermott Technology ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Bruce W. Arey referred in his request to find a vacuum leak in his JSM840 to 'radioactive particles' inside the chamber of his SEM. I wonder if he could be more specific, because I feel if we start calling a SEM as a 'radioactive system' many safety officers will have a field day. I have been working with various SEMs since 1968 and never felt that I might be bombarded by radioactive particles, even when I opened the chamber (e.g. ETEC) of the SEM.
Hans Brinkies Senior Lecturer Swinburne, University of Technology School of Engineering and Science P.O.Box 218 - Hawthorn - Vic -3122 - Australia Phone: +61 3 9214 8657 Fax: +61 3 9214 8264 Email: Hbrinkies-at-swin.edu.au
Dear Friends, For a long time we used Fab fragments-HRP from BioSys (France) for the immuno EM labelling of proteins with subsequent preembedding. These fragments were the best. However, recently the company cancelled its activity and does not send any more these products. Would you be so kind to tell me what has happened (if you know) and what fragments have the same quality?
The Laboratory of Electron Microscopy is looking for an used critical point drier to obtain it in donation.... We can pay all the costs of shipping and handling For any questions, please contact to me....
best regards.... =================================================== Fernando D. Balducci Laboratorio de Microscopia Electr—nica Facultad de Ingenier’a - Bioingenieria Universidad Nacional de Entre Rios Argentina e-mail: microsc-at-fi.uner.edu.ar tel: 54 43 975100 fax: 54 43 975077
I have done this on a JEOL 4000 but not a Zeiss. I used a Maglev TMP (Seiko Seiki) and an antivibration bellows to couple the pump because I was afraid of vibration degrading the 0.25nm resolution. I need not have worried, even with the antivibration bellows shorted out I was OK. Check that the TMP you choose does not give out any magnetic fields when running. If vibration is a problem then the Balzers (Pfieiffer) antivibration bellows that has a large worm drive clip around them can be tuned (by tightening the clip) to avoid the instrument resonant frequency.
Good luck, Ron
---------------------- Mr. R.C. Doole Department of Materials, University of Oxford. Parks Road, Oxford. OX1 3PH. UK. Phone +44 (0) 1865 273701 Fax +44 (0) 1865 283333 ron.doole-at-materials.ox.ac.uk
On Tue, 13 Jun 2000 09:17:46 +0200 Per =?iso-8859-1?Q?H=F6rstedt?= {per.horstedt-at-pathol.umu.se} 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, } one of our TEM's , a Zeiss EM 109, is equipped with an ion getter hi-vac } pump. We are now planning to install a turbopump including all the piping } and valves that are necessary. The reason for this is that the high vacuum } is not good enough ( only in low E-6 area ) and the ion pump has to work } too hard and the life-time of the electrodes becomes very short. Also the } housing is clogged with trapped gas molecules and has to be regenerated far } too often. } Technically and electronically this exchange is fairly easily done, but my } question is if anyone has done it and if so, what's the experience? } Yours sincerely } } Per Hörstedt } Department of Medical Biosciences } Pathology } Unit for Electron Microscopy } University of Umeĺ } S-90187 Umeĺ } Sweden } } phone int-46-90-7851541 } fax int-46-90-7851215
After many years of staining grids with uranyl acetate and lead citrate, we have begun to see a needle like or shard precipitate, (about 1/2 inch long at 100,000x's; resembles the lead precipitate on page 469 of Electron Microscopy second edition John Bozzola and Lonnie Russell). We have been using a 2.5% aqueous uranyl acetate and Reynold's lead citrate (filtered through a 2 micron filter) for the past 4 years or so with no problems. I have stained the grids with just UA and can see no precipitate and have stained grids with just the lead citrate and still not see the precipitate. I have also checked the water and cannot still see the precipitate. However, when I stain with UA followed by lead citrate it mysteriously reappears much to my dissatisfaction. I have also tried the basic lead citrate and just recently tried Sato's lead stain and had the same problem. I have made up UA from a newly purchased bottle. I have also lessened the staining times from 30 minutes in UA to 7 minutes and from 20 minutes in lead citrate to 5 minutes and the precipitate is less but still there. I have also checked the grids before staining them and cannot see the precipitate. Please help, I grow more grey day by day.
Phoebe J. Doss Manager/Adjunct Instructor Electron Microscope Lab Oklahoma State University
Back in the olden days, when BioRad sold microscopy supplies, they had an epoxy cleaner (to remove epoxies from hands, benches, etc., not made from epoxies). It was blue gunk in a little jar. Does anyone know what happened to this stuff? Or have an alternate?
I've just been digging through catalogues and Ted Pella sells a liquid cleaner - any experience with it?
a student here needs to fix and dehydrate strep mutans on polystyrene petri dishes. Can anyone point me to a good (simple?) protocol? Since we are primarily a materials research lab, we don't have a lot of biological references.
I don't know the Bio-Rad product, but Ladd sold (still sells?) a product called Met-a-terge that gets rid of uncured resins. A little bit goes a long way. I've used it for years.
John Mansfield's post regarding surplus equipment prompts me to make the following observations regarding the transfer of University-owned equipment in the United States. I would imagine that many other countries might have similar policies.
It often seems to come as a surprise to people to discover that the US government won't buy the same piece of equipment twice. What I mean by this is that if a piece of equipment has originally been purchased using federal funds (regardless of who currently holds title), then another institution cannot use federal funds (regardless of the source) to buy the equipment from the first owner.
To use the specific example of John's equipment: suppose it was bought originally with, say, an NSF grant, and I find that I could make use of it now. In order to do that I would need to find a non-US-government source of money, as I could not use even the income from my facility operation (which is regarded by the accountants as government money, as it originates predominantly from government research grants).
The logic of this policy, of course, is quite inescapable, however unpalatable it may be to the present owners of the equipment.
The policy only covers the cost of purchasing the equipment, by the way. If, to use the same example, John were to give me his surplus equipment, it would be perfectly acceptable for me to use federal funds to pay for the packing and shipping (and, if appropriate, reinstallation)
Tony Garratt-Reed.
** Anthony J. Garratt-Reed ** MIT Room # 13-1027 ** 77 Massachusetts Avenue ** Cambridge, MA 02139-4307 ** USA ** ** Phone: (+) 1-617-253-4622 ** Fax: (+) 1-617-258-6479 **
I am using a Denton Desk II sputter coater with a Pt target. I have noticed recently that a dark spot has formed in the middle of the target. I have not seen this before. Is this due to impurities in the target, problems with the vacuum, contamination from outgassing samples, or something else? Everything appears to be running fine and sample types have not changed. Do I clean it or just leave it alone? What is it telling me (if the coater could talk)?
Thanks in advance for all your expertise.
David BG Rose WL Gore and Associates 297 BLue Ball Road Elkton, MD 21921 410-506-2958
We recently had a rash of horrible precipitate problems that we couldn't seem to trace. Our staining procedures are similar to yours. We made up fresh stains, changed all our syringe filters, used every precaution we could think of.
Then we had the water system checked. Our in-line reverse-osmosis, deionization system had become a mess, although we had assumed (there's that word!) that the company we leased it from was maintaining it properly. Turns out that they thought we owned the system, while in fact we only rented it and paid them to service it.
Anyway, to keep it short, we purchased a Millipore bench-top, low-volume water-polishing unit and used our old water to feed it (after getting the thing serviced properly). Our precipitates disappeared and have not yet reappeared.
For what it's worth.
(No financial interest in Millipore, etc.)
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
-----Original Message----- } From: Phoebe J Doss [mailto:pjdoss-at-okstate.edu] Sent: Tuesday, June 13, 2000 9:33 AM To: microscopy-at-sparc5.microscopy.com
Hello:
After many years of staining grids with uranyl acetate and lead citrate, we have begun to see a needle like or shard precipitate, (about 1/2 inch long at 100,000x's; resembles the lead precipitate on page 469 of Electron Microscopy second edition John Bozzola and Lonnie Russell). We have been using a 2.5% aqueous uranyl acetate and Reynold's lead citrate (filtered through a 2 micron filter) for the past 4 years or so with no problems. I have stained the grids with just UA and can see no precipitate and have stained grids with just the lead citrate and still not see the precipitate. I have also checked the water and cannot still see the precipitate. However, when I stain with UA followed by lead citrate it mysteriously reappears much to my dissatisfaction. I have also tried the basic lead citrate and just recently tried Sato's lead stain and had the same problem. I have made up UA from a newly purchased bottle. I have also lessened the staining times from 30 minutes in UA to 7 minutes and from 20 minutes in lead citrate to 5 minutes and the precipitate is less but still there. I have also checked the grids before staining them and cannot see the precipitate. Please help, I grow more grey day by day.
Phoebe J. Doss Manager/Adjunct Instructor Electron Microscope Lab Oklahoma State University
Transmission Electron Microscopist (TEM) / Engineer
United Technologies Corporation is seeking an engineer to fill the TEM operator/engineer position at the United Technologies Research Center in East Hartford, CT. This position will provide support to the United Technologies Corporation Business Units including Pratt & Whitney, Carrier, Sikorsky Aircraft, Hamilton Sundstrand, Otis Elevator, and International Fuel Cells. The TEM operator will be responsible for the dailyoperations of the TEM laboratory; including preparation of TEM samples using various techniques such as dimpling, ion milling, jet polishing, microtoming, and replication. Project duties include conducting failure analyses, characterization of surface coatings, and analysis of advanced metal and ceramic materials. The candidate should have experience with both TEM sample preparation and conventional TEM operation. Good communication and interpersonal skills are essential. The ability to recognize fracture modes and origins of fractures is desired. Experience with Scanning Electron Microscopy (SEM) is a plus.
Qualified candidates will have a BS in Materials Science or equivalent, with a minimum of 2 years TEM experience. U. S. citizenship or permanent residency is required.
Please visit our web site at www.utrc.utc.com, and send your resume to Employment Opportunities, Code MATS-2050-9049, 411 Silver Lane, East Hartford, CT 06118 or e-mail employment-at-utrc.utc.com. United Technologies Corporation is an equal opportunity employer.
} } Back in the olden days, when BioRad sold microscopy supplies, they had an } epoxy cleaner (to remove epoxies from hands, benches, etc., not made from } epoxies). It was blue gunk in a little jar. Does anyone know what happened } to this stuff? Or have an alternate? } } I've just been digging through catalogues and Ted Pella sells a liquid } cleaner - any experience with it? }
I seem to remember that soaking in N,N-dimethyl formamide dissolves epoxy, you might care to beg a little from a freiendly chemistry dept and try it, it's not very expensive. It has a moderately offensive smeel, though.
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
Yes, Ladd stills sells Met-A-Terge (catalog #13045). Please check our web site http://www.laddresearch.com for more information or contact me off line.
JD Arnott
Disclaimer: As stated above, Ladd sell Met-A-Terge and thus has a commercial interest in this thread.
LADD RESEARCH 131 Dorset Lane Williston, VT 05495 USA
TEL 1-800-451-3406 (US) or 1-802-878-6711 (anywhere) FAX 1-802-878-8074 e-mail ladres-at-worldnet.att.net
ALL NEW WEB SITE UP NOW AT OUR NEW URL:
http://www.laddresearch.com
Beverly_E_Maleeff-at-sbphrd.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 } -----------------------------------------------------------------------.} } Tamara: } } I don't know the Bio-Rad product, but Ladd sold (still sells?) a product called } Met-a-terge that gets rid of uncured resins. A little bit goes a long way. } I've used it for years. } } Hope this helps. } } Regards, } Bev Maleeff } SmithKline Beecham Pharmaceuticals
Hi Tamara, I have used the Ted Pella Epoxy Hand Cleaner and it works well. It will remove epoxy from hands and also glassware. Jo Dee Fish
PS I am not affiliated with Ted Pella, just love their products!
Tamara Howard 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 } -----------------------------------------------------------------------. } } Back in the olden days, when BioRad sold microscopy supplies, they had an } epoxy cleaner (to remove epoxies from hands, benches, etc., not made from } epoxies). It was blue gunk in a little jar. Does anyone know what happened } to this stuff? Or have an alternate? } } I've just been digging through catalogues and Ted Pella sells a liquid } cleaner - any experience with it? } } Thanks! } } Tamara Howard } CSHL
-- Jo Dee Fish Electron Microscopy Assistant Cell Analysis Facility The Burnham Institute 10901 N. Torrey Pines Rd. La Jolla, CA 92037 (858)646-3100 ext. 3620
My husband uses epoxy on a sailboat he's building. He cleans everything up... hands, spills, etc... with plain old vinegar. We go through a LOT of vinegar. If it's dried, then soak acetone on it until it softens, then use vinegar (or 5% acetic acid) to mop up the residues.
connie m
At 10:31 AM 06/13/2000 -0400, Tamara Howard wrote: } Back in the olden days, when BioRad sold microscopy supplies, they had an } epoxy cleaner (to remove epoxies from hands, benches, etc., not made from } epoxies). It was blue gunk in a little jar. Does anyone know what happened } to this stuff? Or have an alternate? } } I've just been digging through catalogues and Ted Pella sells a liquid } cleaner - any experience with it? } } Thanks! } } Tamara Howard } CSHL } } } Connie McManus Veterinary Diagnostics Lab Utah State University Logan, UT USA
} To use the specific example of John's equipment: suppose } it was bought originally with, say, an NSF grant, and } I find that I could make use of it now. In order to do } that I would need to find a non-US-government source } of money, as I could not use even the income from my } facility operation (which is regarded by the accountants } as government money, as it originates } predominantly from government research grants).
I would assume it can even get messy if, at least some, of my facility's income had come from outside sources. I would imagine the accountants will first assume it is government $$ ... in which case I would have to show I had taken in an equivelent in outside $$ ... but over what time frame? ... this fiscal? ... last 10 years?
cheerios, =shAf= :o)
{} /\ {\/} /\ {\/} /\ {\/} /\ cogito, ergo zZOooOM /\ {\/} /\ {\/} /\ {\/} /\ {} Michael Shaffer, R.A. - mshaf-at-darkwing.uoregon.edu Geological Science's Electron Probe Facility - University of Oregon http://epmalab.uoregon.edu/
I use methylene dichloride (dimethyl chloride) to dissolve most any epoxy.
I have 3-5 gallons of it. I do not know if it is still generally available.
gg
At 07:31 AM 6/13/00, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
The listserver is at MSA listserver {Microscopy-at-sparc5.microscopy.com} Just sent email say, "Please subscribe". It is fairly active but is only about 1/2 relevant as most data is biology oriented.
Earl JCNABITY-at-aol.com wrote:
} Dear Earl, } } Could you tell me how to subscribe to the MSA listserver? Greg talked highly } of it and I'm thinking I will set up another e-mail account to use for it. } Since it is pretty active, I didn't want all the messages going in with my } normal e-mail, but a using a separate account will resolve that issue. } } Joe
Our EM Unit has a Zeiss EM109 which uses 70mm roll film - Agfa Scientia 23D56. Manufacture of this film ceased quite a while back. We are trying to locate unused stock of this film for our usage. If you have any surplus stock for sale please contact me.
Thanks Mohamed
****************************** M. A. Jaffer Electron Microscope Unit R. W. James Building University of Cape Town Private Bag Rondebosch, 7701 South Africa
At 06:09 PM 06/13/2000 -0700, Don Hammer wrote: } Stuff is cheap too and if there is any left after the zillions of home uses, } great on salads!!!!
yeah, especially the used stuff........ eeeeuewwwwwwww! *G*
connie m } } Don Hammer, Retired Guy } ----- Original Message ----- } From: Connie McManus {conmac-at-cc.usu.edu} } To: Tamara Howard {howard-at-cshl.org} ; Microscopy Listserver } {Microscopy-at-sparc5.microscopy.com} ; Histology listserver } {histonet-at-pathology.swmed.edu} } Sent: Tuesday, June 13, 2000 2:29 PM } Subject: Re: Epoxy cleaner? } } } } My husband uses epoxy on a sailboat he's building. He cleans everything } } up... hands, spills, etc... with plain old vinegar. We go through a LOT } of } } vinegar. If it's dried, then soak acetone on it until it softens, then } use } } vinegar (or 5% acetic acid) to mop up the residues. } } } } connie m } } } } At 10:31 AM 06/13/2000 -0400, Tamara Howard wrote: } } } Back in the olden days, when BioRad sold microscopy supplies, they had an } } } epoxy cleaner (to remove epoxies from hands, benches, etc., not made from } } } epoxies). It was blue gunk in a little jar. Does anyone know what } happened } } } to this stuff? Or have an alternate? } } } } } } I've just been digging through catalogues and Ted Pella sells a liquid } } } cleaner - any experience with it? } } } } } } Thanks! } } } } } } Tamara Howard } } } CSHL } } } } } } } } } } } Connie McManus } } Veterinary Diagnostics Lab } } Utah State University } } Logan, UT } } USA } } } } } } Connie McManus Veterinary Diagnostics Lab Utah State University Logan, UT USA
Hi to all, Does anybody have some information on a possible relationship between lipid droplets and wax production in plant cells. Are there some evidences that lipid droplets could actually be storage sites of wax precursors ? I looked for that in literature but found nothing... Thus : HELP ! References will be welcome Thanks in advance for answering Bye Pascal
""""""""' ( O)(o ) --------------------0000--------------0000---------- Pascal VEYS Laboratory of Plant Biology Catholic University of Louvain Place Croix du Sud 5 (bte 14) B 1348 Louvain-la-Neuve Belgium Phone : 0032 10473004 Fax : 0032 10473471 Email : Veys-at-bota.ucl.ac.be ---------------------ooooO-----------Ooooo-------- ( ) (_)(_) ( ) ) ( ) ( (_) (_)
Gary Gaugler wrote: =============================================================== I use methylene dichloride (dimethyl chloride) to dissolve most any epoxy. I have 3-5 gallons of it. I do not know if it is still generally available. ================================================================ If we are talking about methylene dichloride, or dichloromethane, CAS # 75- 09-2, this is a pretty bad actor, and is on the list of Prop. 65 chemicals for the State of California as being cancer causing. Chemicals on the Prop . 65 list are so highly restricted that in some organizations, they are allowed in only with the approval of top management.
But I do have a question: I always thought that most epoxies, certainly the ones used in microscopy, ended up being three dimensionally crosslinked intractable solids. The only way such a material is going to be "dissolved" is for chemical bonds to be broken. And yet, I don't see how chemically, methylene dichloride is going to be breaking chemical bonds. Or the same comment for some of the other materials mentioned. These materials might plasticize (e.g. soften) an epoxy and aid in its removal from a surface, but do any of these really "dissolve" a three dimensionally crosslinked epoxy system?
I am very interested in this topic because we believe that at least in terms of getting epoxy out of the "nooks and crannies" of a non-smooth surface, an oxygen plasma is needed. But perhaps we are wrong about that, that is why I ask the question.
Disclaimer: SPI Supplies manufactures the Plasma Prep™ II plasma etcher and has an interest in seeing more applications for plasma etching.
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.spi.cc ######################## ============================================
David, I had the same problem with a gold target-a dark spot in the middle. Also, what was being sputtered on my samples was not gold. I cleaned the target with acetone and it has been working fine since. Joyce Craig Chicago State University
We are working with Leishmania major, a parasite that is incorporated into macrophages. We have had good success with fixation of lymph nodes infected with Leishmania. We have not been as happy with the results of fixation of cell cultures of infected bone marrow macrophages. The membranes of the Leishmania and of the internal compartments within the macrophages that hold the Leishmania are well fixed, but the external cell membranes of the macrophages are somewhat discontiuous. We have not done cell cultures before. Is this a problem to be expected? We are fixing with 2+2 glutaraldehyde/paraformaldehyde in 0.1 M phosphate buffer, post-fixing with 2% buffered Osmium, dehydrating with ethanol and propylene oxide, then embedding in epoxy. We have shortened all times compared to those we use with tissue samples. Joyce Craig Chicago State University
I am looking for a non-fluorescent plastic coverslip that allows confocal laser microscopy and subsequent sectioning for Transmission Electron Microscopy. I will greatly appreciate your suggestions.
Sincerely, Karthi Subramanian Department of Microbiology University of Guelph Guelph Ontario N1G 2W1 Phone: (519)824-4120 ext.8904 Fax:(519)837-1802
I perform stereo measurements only occasionally, so I prefer to save money on specialized equipment or software. All I use is just freeware program ImageTool (good for on-screen stereo pair measurements) and Excel (not bad for calculations).
Of course, for a big project it's better to buy a software.
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: John J. Bozzola [mailto:bozzola-at-siu.edu] } Sent: Thursday, June 08, 2000 11:13 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: Stereometry by computer } } } -------------------------------------------------------------- } ---------- } 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 } } -------------------------------------------------------------- } ---------. } } } A user of our facility is interested in making height measurements } from specimens viewed in the SEM. I am aware of the conventional way } of doing this: stereo pairs and optical viewer (with stereometer } parallax corrections). Is there a more modern (computerized) way of } doing this, say with anaglyphs? } } Thank you. } } John B. } } } #################################################################### } John J. Bozzola, Ph.D., Director } Micro-Imaging and Analysis Center } 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 } Web: http://www.siu.edu/departments/shops/cem.html } #################################################################### }
I need to label Si-OH groups with something that will show up in TEM. If it were proteins or sugars, I'd immuno- or lectin label. Any ideas how I can get gold or ferritin or whatever onto these groups?
Mahalo! 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* ****************************************************************************
} Hello, } } Can anyone recommend a supplier for uranyl acetate? } Also, does anyone out there have experience with Sigma's Lowicryl kit? } If possible, please respond to me directly at: mpilgrim-at-mendelbio.com } } Many thanks, } Marsha
Dear Marsha,
We at Ladd Research, and most of the other supply companies, can sell you this. In our case it is catalog # 23620 and more information can be found on our web site, http://www.laddresearch.com
John Arnott --
LADD RESEARCH 131 Dorset Lane Williston, VT 05495 USA
TEL 1-800-451-3406 (US) or 1-802-878-6711 (anywhere) FAX 1-802-878-8074 e-mail ladres-at-worldnet.att.net
Help - A faculty member's plant tissue is autofluorescing. She is using aniline blue to look at pollen tubes (through the styles). She would like to reduce the background fluorescence. I gave her a copy of a borohyride reference from a '97 listserv posting. Can anyone recommend a fixation that will decrease or eliminate the autofluorescence? Any thoughts on this matter would be greatly appreciated. thanks, Beth
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
It took me a few days to find this protocol from my collegue Lucinda Swatzell. It sounded intriguing. Although I've not tried it myself yet, she's used it with success.
I've copied the following out of her e-mail. Things that need to be kept in mind are that she polymerizes in a vacuum oven, she's refering to plant seedlings, and that I don't have any financial interest that I know of (i.e. I haven't checked my mutual fund prospectus) in Rubbermaid, Inc.
Here it is:
"There is a way to flat embed with LR White: This works great for me when I am keeping them on their agar blocks and maintaining orientation, but it should also work for regular seedlings to keep them flat instead of crooked in the bottom of the capsules. Use rubbermaid ice cube trays. Place the specimens in the flat bottoms of the tray. cover with about 1/4 in of resin. In each well, place another well that has been cut from it's tray. The single loose wells will nestle down into the ice cub tray on top of the resin. Because rubbermaid is dishwasher safe it will take the heat, but get soft enough to snuggle in tightly and keep out oxygen. When you pump the vaccuum the extra resin also snakes up into the cracks, so that you get a seal. Now the thing that is scary: will loose seedlings just suck up into the cracks? I haven't tried them."
Heather A. Owen, Director Electron Microscope Laboratory Department of Biological Sciences University of Wisconsin - Milwaukee (414)229-6816
I've used it often to unpackage epoxy microchips. These epoxies are silicone/epoxy and not true epoxy. They are typically called plastic packaged ICs. Whatever. I've used MEC and DiMEC to do this. Some packages had to be oxygen ion blasted open. Over time, the plasma method has been much safer for the operator and the chip. And it can make the removal process much faster. Plasma is the current method of choice.
Whether the "epoxies" talked about here are the same as those for IC packages is likely not to be.
gary g.
At 09:40 AM 6/14/00, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I used to prepare W FIM emitter tips with a 5% NaOH solution and I think the voltage was around 5 to 10 volts ac. When I used to electropolish, the FIM solutions and conditions were frequently similar to the jet polishing solutions.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center Guys Run Rd. (packages) P. O. Box 11472 (letters) Pittsburgh, PA 15238-0472
Walck-at-PPG.com
(412) 820-8651 (office) (412) 820-8161 (fax)
} -----Original Message----- } From: Gillian Bond [mailto:gbond-at-nmt.edu] } Sent: Monday, June 12, 2000 7:43 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Jet polishing of tungsten } } } -------------------------------------------------------------- } ---------- } 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 have a student who has been trying to jet polish tungsten } for TEM. She } has tried various concentrations of sodium hydroxide in } water, as well as } 40g trisodium phosphate/250ml water, and 55.8g magnesium } perchlorate/250ml } methanol, at a range of voltages. We have a Fischione jet-polishing } unit. So far, none of the samples has been close to good. Can anyone } help us out here, with past experience or general suggestions? } } Many thanks in advance, } } Gill } } Dr Gillian M. Bond } Department of Materials & Metallurgical Engineering } New Mexico Tech } Socorro, NM 87801 } }
My name is Tim Strovas and I am a graduate student at the U of Washington bioengineering department. I am investigating the halo effect from single myofibrils (from bumblebee muscle tissue) as seen under a phase contrast microscope. Has anyone encountered a previous investigation into this effect and its possible relationship with structured water? Note: The Halo does not appear as ripples that are normally associated with optical microscope artifacts. The halo is single broad band that borders the tissue sample.
My name is Tim Strovas and I am a graduate student at the U of Washington bioengineering department. I am investigating the halo effect from single myofibrils (from bumblebee muscle tissue) as seen under a phase contrast microscope. Has anyone encountered a previous investigation into this effect and its possible relationship with structured water? Note: The Halo does not appear as ripples that are normally associated with optical microscope artifacts. The halo is single broad band that borders the tissue sample.
My name is Tim Strovas and I am a graduate student at the U of Washington bioengineering department. I am investigating the halo effect from single myofibrils (from bumblebee muscle tissue) as seen under a phase contrast microscope. Has anyone encountered a previous investigation into this effect and its possible relationship with structured water? Note: The Halo does not appear as ripples that are normally associated with optical microscope artifacts. The halo is single broad band that borders the tissue sample.
G'day folks, Be very wary of any Good Epoxy Solvent, do not use it to clean epoxy off your skin. Anything that is a good solvent for epoxy will probably be a good solvent for the oils and lipids in/on your skin. These oils and lipids are your protection against epoxy resins entering your body. Remember, all epoxy resins at carcinogenic, soap and water are probably the safest agents to remove epoxies from your skin. Regards JVN
Connie McManus 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 } -----------------------------------------------------------------------. } } At 06:09 PM 06/13/2000 -0700, Don Hammer wrote: } } Stuff is cheap too and if there is any left after the zillions of home uses, } } great on salads!!!! } } yeah, especially the used stuff........ eeeeuewwwwwwww! *G* } } connie m } } } } Don Hammer, Retired Guy } } ----- Original Message ----- } } From: Connie McManus {conmac-at-cc.usu.edu} } } To: Tamara Howard {howard-at-cshl.org} ; Microscopy Listserver } } {Microscopy-at-sparc5.microscopy.com} ; Histology listserver } } {histonet-at-pathology.swmed.edu} } } Sent: Tuesday, June 13, 2000 2:29 PM } } Subject: Re: Epoxy cleaner? } } } } } } } My husband uses epoxy on a sailboat he's building. He cleans everything } } } up... hands, spills, etc... with plain old vinegar. We go through a LOT } } of } } } vinegar. If it's dried, then soak acetone on it until it softens, then } } use } } } vinegar (or 5% acetic acid) to mop up the residues. } } } } } } connie m } } } } } } At 10:31 AM 06/13/2000 -0400, Tamara Howard wrote: } } } } Back in the olden days, when BioRad sold microscopy supplies, they had an } } } } epoxy cleaner (to remove epoxies from hands, benches, etc., not made from } } } } epoxies). It was blue gunk in a little jar. Does anyone know what } } happened } } } } to this stuff? Or have an alternate? } } } } } } } } I've just been digging through catalogues and Ted Pella sells a liquid } } } } cleaner - any experience with it? } } } } } } } } Thanks! } } } } } } } } Tamara Howard } } } } CSHL } } } } } } } } } } } } } } } Connie McManus } } } Veterinary Diagnostics Lab } } } Utah State University } } } Logan, UT } } } USA } } } } } } } } } } } Connie McManus } Veterinary Diagnostics Lab } Utah State University } Logan, UT } USA
-- **************************************************** John V Nailon Operations Manager Centre for Microscopy and Microanalysis The University of Queensland St. Lucia Queensland 4072 Phone: +61-7-3365-4214 Fax: +61-7-3365-4422 WWW: http://www.uq.edu.au/nanoworld/allstaff.html#Nailon ****************************************************
I suggest you copy this message to the plant surfaces mail list
To join, send the command join plant-surfaces firstname lastname to: mailbase-at-mailbase.ac.uk "Firstname" can be one or more names or initials. The last word in this command will be interpreted as the last name. The email address will be extracted automatically from the message.
Chris Jeffree
Date sent: Wed, 14 Jun 2000 16:18:08 +0100 To: microscopy-at-sparc5.microscopy.com } From: veys-at-bota.ucl.ac.be (Pascal Veys)
Hello Tina:
I would approach your problem by either modifying the Si-OH groups with a hapten and detecting with antibody- or streptavidin-gold, or converting them to amines or thiols then labeling with a gold labeling reagent (disclaimer - we make gold labeling reagents). You could introduce amino- groups at the Si-OH groups using a silylating reagent such as 3-{Tris[2-(2-methoxyethoxy)ethoxy]silyl}propylamine or 3-[Tris(trimethylsiloxy)silyl]propylamine (both from Fluka), then either biotinylate with NHS-biotin and detect with streptavidin-gold, or label the amines with Mono-Sulfo-NHS-Nanogold.
I have not actually tried this, and since I don't know what types of samples you are looking at, it's difficult to say what else in them might affect the reaction. If there are already other primary amines in your sample, they need to be blocked first.
If you would like other ideas, a text on solid-phase oligo- or peptide synthesis might be another good starting point - the chemistry used to functionalize the beads used in these systems may also be transferable to your situation.
Hope this is helpful,
Rick Powell
} } Oh wise and helpful microscopists- } } I need to label Si-OH groups with something that will show up in TEM. If } it were proteins or sugars, I'd immuno- or lectin label. Any ideas how I } can get gold or ferritin or whatever onto these groups? } } Mahalo! } 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* } ****************************************************************************ť
John Nailon is making a very good argument for using gloves or working very cleanly. All epoxies I understand are "somewhat" carcinogenic. The much quoted John Luft, years ago advised me that photographic fixer (sodium thiosulphate) solution, chemically changed epoxies so they would not be carcinogenic. If he was right, then first washing any body parts contaminated by epoxy resin in photographic fixer should avert the worse. Those fixers do not dissolve or clean epoxies. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Thursday, June 15, 2000 11:36 AM, John Nailon [SMTP:mmjnailo-at-dingo.cc.uq.edu.au] wrote: } } G'day folks, } Be very wary of any Good Epoxy Solvent, do not use it to clean epoxy off } your skin. Anything that is a good solvent for epoxy will probably be a } good solvent for the oils and lipids in/on your skin. These oils and } lipids are your protection against epoxy resins entering your body. } Remember, all epoxy resins at carcinogenic, soap and water are probably } the safest agents to remove epoxies from your skin. } Regards } JVN } } Connie McManus 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 } } -----------------------------------------------------------------------. } } } } At 06:09 PM 06/13/2000 -0700, Don Hammer wrote: } } } Stuff is cheap too and if there is any left after the zillions of home } } } uses, } } } great on salads!!!! } } } } yeah, especially the used stuff........ eeeeuewwwwwwww! *G* } } } } connie m } } } } } } Don Hammer, Retired Guy } } } ----- Original Message ----- } } } From: Connie McManus {conmac-at-cc.usu.edu} } } } To: Tamara Howard {howard-at-cshl.org} ; Microscopy Listserver } } } {Microscopy-at-sparc5.microscopy.com} ; Histology listserver } } } {histonet-at-pathology.swmed.edu} } } } Sent: Tuesday, June 13, 2000 2:29 PM } } } Subject: Re: Epoxy cleaner? } } } } } } } } } } My husband uses epoxy on a sailboat he's building. He cleans everything } } } } up... hands, spills, etc... with plain old vinegar. We go through a LOT } } } of } } } } vinegar. If it's dried, then soak acetone on it until it softens, then } } } use } } } } vinegar (or 5% acetic acid) to mop up the residues. } } } } } } } } connie m } } } } } } } } At 10:31 AM 06/13/2000 -0400, Tamara Howard wrote: } } } } } Back in the olden days, when BioRad sold microscopy supplies, they had } } } } } an } } } } } epoxy cleaner (to remove epoxies from hands, benches, etc., not made } } } } } from } } } } } epoxies). It was blue gunk in a little jar. Does anyone know what } } } happened } } } } } to this stuff? Or have an alternate? } } } } } } } } } } I've just been digging through catalogues and Ted Pella sells a liquid } } } } } cleaner - any experience with it? } } } } } } } } } } Thanks! } } } } } } } } } } Tamara Howard } } } } } CSHL } } } } } } } } } } } } } } } } } } } Connie McManus } } } } Veterinary Diagnostics Lab } } } } Utah State University } } } } Logan, UT } } } } USA } } } } } } } } } } } } } } } } Connie McManus } } Veterinary Diagnostics Lab } } Utah State University } } Logan, UT } } USA } } -- } **************************************************** } John V Nailon } Operations Manager } Centre for Microscopy and Microanalysis } The University of Queensland } St. Lucia Queensland 4072 } Phone: +61-7-3365-4214 } Fax: +61-7-3365-4422 } WWW: http://www.uq.edu.au/nanoworld/allstaff.html#Nailon } ****************************************************
I needed to have a cross sectional view into an adhesive layer so that I could count the layers, if possible. Since the adhesive acts like a highly viscous liquid, polishing it is out of the question. Instead, I submerged the film into liquid Nitrogen and cut it quickly w/ a scissors. The sound of the cut was more like a breaking sound. When I examined the cross sectional surface I observed a brain like surface. There were columns with a highly consistent diameter and orientation. It looked crystalline. There was zero evidence of material smearing that one would expect if one cut a material. Is the proper interpretation that these columns existed before the fracture and that the fracture occurred along the boundaries? Or is the cross sectional surface generated by a rippled distortion of an highly viscous liquid? The regularity of the surface seems to make the latter interpretation unlikely.
I would be interested in any opinions on the interpretation of the image or alternative means of cross sectioning the adhesive layer.
I could e-mail an image to anyone who is interested.
We haven't had a need to unpackage 'plastic' encapsulated IC's for a while but may have a need to do so soon (preferrably not impairing functionality?!). I am told that the last person to do this here (now retired) dripped fuming sulfuric acid on the plastic and used frequent water rinses. We have a plasma etcher but I was afraid it would take forever to get through the plastic.
Any hints and suggestions would be greatly appreciated.
Diane Ciaburri Senior Materials Engineer General Dynamics 100 Plastics Ave. Pittsfield MA 01210
I've used it often to unpackage epoxy microchips. These epoxies are silicone/epoxy and not true epoxy. They are typically called plastic packaged ICs. Whatever. I've used MEC and DiMEC to do this. Some packages had to be oxygen ion blasted open. Over time, the plasma method has been much safer for the operator and the chip. And it can make the removal process much faster. Plasma is the current method of choice.
Whether the "epoxies" talked about here are the same as those for IC packages is likely not to be.
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Hi, All:
This one is not really related to microscopy. But since I am a TEM guy, I hope I can get some help here.
Does anybody have some information about MgAl2O4 as a substrate for perovskite films? I know it is not often used, so I am worndering if it has a major disadvantage so that nobody is using it. Any references would be welcome.
I have a control PC board for the E5200 sputter coater. This is the model with an Intel single chip MPU on one end and a 4-conductor socket on the other end. The board uses a VME connector for main interface.
Coater is trashed. Board is OK. If anybody can use the board, first request gets it.
Beth, more details are needed on how the tissue was processed before viewing. I have used aniline blue to view pollen tubes in style of Salicornia virginica which had ben fixed in Nawashin's fixative. Have also viewed tubes of Melilotus which had simply been preserved in 70% EtOH. If one uses glut as a fixative, it fluoresces so you won't be able to distinguish the PT from everything else. Mary Pfauth
Can't comment on sulfuric, but I have used red fuming nitric at near boiling temperature. Apply acid, let react. Flush witn more acid, let react, etc.
Woody White
The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Gary and others,
We haven't had a need to unpackage 'plastic' encapsulated IC's for a while but may have a need to do so soon (preferrably not impairing functionality?!). I am told that the last person to do this here (now retired) dripped fuming sulfuric acid on the plastic and used frequent water rinses. We have a plasma etcher but I was afraid it would take forever to get through the plastic.
Any hints and suggestions would be greatly appreciated.
Diane Ciaburri Senior Materials Engineer General Dynamics 100 Plastics Ave. Pittsfield MA 01210
I've used it often to unpackage epoxy microchips. These epoxies are silicone/epoxy and not true epoxy. They are typically called plastic packaged ICs. Whatever. I've used MEC and DiMEC to do this. Some packages had to be oxygen ion blasted open. Over time, the plasma method has been much safer for the operator and the chip. And it can make the removal process much faster. Plasma is the current method of choice.
Whether the "epoxies" talked about here are the same as those for IC packages is likely not to be.
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Hello,
Does anyone know the shear modulus for LaAlO3?
Thanks
Yan Xin ======================================= Yan Xin Magnet Science & Technology National High Magnetic Field Laboratory Florida State University 1800 E. Paul Dirac Drive Tallahassee, FL 32310 Tel: (850) 644 1529 Fax: (850) 644 0867 ========================================
The ion beam approach works well. I have not used it recently on finer pitch ICs. With as-built feature sizes of 2-4u, it is fine. It will stop at the passivation and leave the Al bond wires intact. The resulting package looks like it has a V-shaped pit in it (which it does). The extent of the pit depends on the size of the die and if you want to blast down to the lead frame or substrate.
I have not done this on finer pitch devices. I would be a bit skeptical about these mostly because of the smaller bond pads. The etching would still stop at the passivation.
There are numerous places in Silicon Valley that do this on an outsource basis. Typical costs are about $75 per IC. I can get some contacts for you if you'd like.
gary g.
At 06:55 AM 6/15/00, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
MME is currently conducting research through Microscopy & Analysis regarding the impact of the internet on microscopy and imaging facilities. If you have not yet faxed back your responses, we'd appreciate your participation. The questionnaire is in the center of the May issue of M&A.
Results of this survey will be reported in a Fall issue of Microscopy & Analysis; specific information on the impact of the internet will be presented along with data collected from other recent MME surveys and reports from other meetings in the "Microbrew" column in the July issue of Advanced Imaging.
Many thanks.
Best regards, Barbara Foster Microscopy/Marketing & Education 125 Paridon Street, Suite 102 Springfield, MA 01118-2130 PH: 413-746-6931 FX: 413-746-9311 email:mme-at-map.com
To the wealth of knowledge on the Microscopy list server,
I have a question about Osmium fixation..Basically I was curious to know if there are any references to Osmium fixation at room temperature? Everything I have seen so far only talks about fixation for 2 hours in the refrigerator at 4 degrees...
Are there any drawback or problems that can occur if tissues specifically Kidney and muscle would or could have? i.e. precipitation etc... etc....
} } We haven't had a need to unpackage 'plastic' encapsulated IC's for a while but } may have a need to do so soon (preferrably not impairing functionality?!). I am } told that the last person to do this here (now retired) dripped fuming sulfuric } acid on the plastic and used frequent water rinses. We have a plasma etcher but } I was afraid it would take forever to get through the plastic. } } Any hints and suggestions would be greatly appreciated. } } Diane Ciaburri } Senior Materials Engineer } General Dynamics } 100 Plastics Ave. } Pittsfield MA 01210
Diane, Yes, hot fuming sulfuric and/or hot fuming nitric are used as standard procedures for removing plastic from IC's. The process is not quite that simple. For example, water rinses will almost certainly etch the bond pads on the IC and thus removing connection to the outside world. Additionally, the plastic contains fire retardants which some regions don't like being washed down the drain. There is more detailed help through EDFAS.org (one of ASM's branches). B&G International sells a very safe, effective etcher which performs decapsulation automatically in minutes.
I have no association with B&G International.
David Saxon Analytical Microscope Services 11826 Reservoir Rd. E. Puyallup, WA 98374 253-848-7701 voice & fax email: info-at-analyticalmicroscope.com website: www.analyticalmicroscope.com
I am looking for information about an analysis software package by the name of ONCOR. Does any have an adress for the company.... which may not exist anymore?? Thanks Blystone in Texas
Robert V. Blystone, PH.D. Professor of Biology Trinity University San Antonio, Texas 78212 rblyston-at-trinity.edu 210-999-7243 FAX 210-999-7229
Being unable to afford a digital camera for my TEM. I'm wondering if the next best option is to get a high end scanner to scan in negatives and then print them on a decent printer. Any advice regarding this idea and brands of scanners and printers that are useful? Also, what image analysis systems are user friendly?
Dr. Thomas P. Bonner Department of Biological Sciences SUNY at Brockport Brockport, NY 14420
I'm looking for advice on embedding bovine oocytes (~100 micron diameter). I'd like to embed them in araldite for probing with labelled lectins as this seems to be fairly well established in the literature. The hangup is this: we are using fairly large plastic cassettes and are having problems losing the oocytes within the volume of araldite. Does anyone know of a way around this? Is there something we can pre-embed the oocytes in to make a smaller chip that we can then embed in the larger block (that is, of course, compatible with polymerizing / clearing the araldite? Or does anyone know of an altogether different method for oocyte embedding that is more effective? Thanks in advance!
--Carrie Golash
Carrie Golash John O. Almquist Research Center Penn State University University Park, PA 16802 W: (814) 865-5896 H: (814) 692-7926 http://www.das.psu.edu/dbrc/dbrc.htm
Our materials science microscopy lab is in need of a used atomic force microscope. No specific model or make. All reasonable offers will be considered. Please respond directly to Don Kierstead at or call 330-794-6600. Any help in this effort would be greatly appreciated.
Dichloromethane and dimethylformamide are relatively effective disrupters of most epoxies but their action is accompanied by great swelling because the polymer becomes engorged with the liquid before any significant solvation takes place. This will destroy wire bonds on an IC.
Fuming (essentially anhydrous) sulfuric acid acts by the completely different process of sulfonating reactive groups that remain on the polymer. The depolymerized and sulfonated byproducts are quite soluble not only in the acid but usually in water as well. The worst thing that you could do in this relatively straightforward process is to wash with water at intervals because this would initiate almost instantaneous corrosion. It would be advisable for a chemist, as someone trained in the handling of reactive materials, to carry this out or at least to establish procedures and train others with less experience. The action of sulfuric acid in this regard is quite different than that of nitric. Nearly anhydrous nitric acid (completely anhydrous is extremely difficult to prepare) is a very powerful oxidizer and could lead to unstable, dangerous byproducts whereas the sulfonates resulting from the sulfuric acid reaction are relatively stable. Water must, of course, be prevented from splashing into any concentrated acid, especially sulfuric.
A very strong acid such as sulfuric behaves completely differently in the absence of water. Since most acids are highly hygroscopic and are sold as water solutions, most people do not observe this other side of their behavior. Without water to create an ionized electrolyte, corrosion of metals will not take place. I have de-encapsulated ICs for failure analysis in 200 degree sulfuric acid and been able to operate the IC without replacing the .001" aluminum wirebonds that it came with. I recall one instance where our company built prototype hybrid microelectronic circuits out of such de-encapsulated ICs when their supplier was late getting a new design on the market and the only ones available were already encapsulated.
The key is to realize that water must be excluded until the sulfonating acid has been completely rinsed away by a non-aqueous liquid. As Mr. Saxon said, there are simple and safe devices available for doing this operation. However, with proper care and protective gear it can be done in a beaker on a hot plate in a fume hood. A few ml.s of sulfuric acid are heated to drive off water until heavy vapors are observed over the liquid (which may darken during heating due to trace impurities). The IC is carefully lowered into the hot acid and a vigorous reaction ensues with the epoxy almost instantly washing into the solution. After a few seconds the IC is then quickly lifted out and held over a receiving vessel and flooded with a stream of ethanol. Only after this is a final rinse in deionized water carried out, followed by fresh electronic grade ethanol and forced drying in warm air.
The ready made devices which carry out the operation are typically a small bowl with a hinged lid from which air is withdrawn by a gentle vacuum. An inert metal feeder tube leads from a heated reservoir for the sulfuric acid and passes through the wall of the bowl to a position where the encapsulated device is secured. When the lid is closed and the slight vacuum applied, the hot acid is pulled into the bowl over the device. It is somewhat self-limiting in that, if the lid is opened, there is no driving force to bring more acid into the container. Naturally, the vacuum source needs to be protected by a trap and all waste products properly handled no matter how the procedure is carried out.
John Twilley Conservation Scientist (formerly, Manager of the Reliability Analysis Center, Teledyne Microelectronics)
DAVID I SAXON 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 haven't had a need to unpackage 'plastic' encapsulated IC's for a while but } } may have a need to do so soon (preferrably not impairing functionality?!). I am } } told that the last person to do this here (now retired) dripped fuming sulfuric } } acid on the plastic and used frequent water rinses. We have a plasma etcher but } } I was afraid it would take forever to get through the plastic. } } } } Any hints and suggestions would be greatly appreciated. } } } } Diane Ciaburri } } Senior Materials Engineer } } General Dynamics } } 100 Plastics Ave. } } Pittsfield MA 01210 } } Diane, } Yes, hot fuming sulfuric and/or hot fuming nitric are used as standard } procedures for removing plastic from IC's. The process is not quite that } simple. For example, water rinses will almost certainly etch the bond pads } on the IC and thus removing connection to the outside world. Additionally, } the plastic contains fire retardants which some regions don't like being } washed down the drain. There is more detailed help through EDFAS.org (one } of ASM's branches). B&G International sells a very safe, effective etcher } which performs decapsulation automatically in minutes. } } I have no association with B&G International. } } David Saxon } Analytical Microscope Services } 11826 Reservoir Rd. E. } Puyallup, WA 98374 } 253-848-7701 voice & fax } email: info-at-analyticalmicroscope.com } website: www.analyticalmicroscope.com
We often work with clients who wish to look inside materials. Firstly the SEM is very clever it will tell you if a material is cut with a blade or a knife or scissors!
The only way to see the true internal structure of a material is to fracture it. Drop the material into LN2 wait until the bubbles stop and then take it out and using heavy duty tweezers crack it.
If a material (like hair and some polymer fibres) will not crack you need to support them in some way to make them crack. We use a water based carbon solution and two SEM stubs. Glue the two stubs together with the water soluble adhesive (try Spi) and then drill two or three small holes through the stubs (about 1mm diameter). Glue the hairs together with the carbon solution and pass then through the holes (messy). When all is dry plunge into LN2. Tap a blade between the two stubs and ALL the material should fracture.
Alternatively, take a fine bore drinking straw and pass the hairs plus carbon solution into the straw. Wait until dry, dump in LN2 and flex the straw to crack it and its contents.
Such fractures of layered materials (e.g. paints) will be best viewed in BSE each "phase" will either be of a different contrast or fracture in a different way. Great fun, try it?
Steve Chapman Senior Consultant Protrain For consultancy and professional training in EM world wide Tel 44+ 1280 814774 Fax 814007 www.emcourses.com
Hi, I want to contact by Email Prof. N.D. Hallam (formerly at Melbourne and LA Tobe - Australia) Does anybody have his contact adress Thanks to all in advance Pascal
""""""""' ( O)(o ) --------------------0000--------------0000---------- Pascal VEYS Laboratory of Plant Biology Catholic University of Louvain Place Croix du Sud 5 (bte 14) B 1348 Louvain-la-Neuve Belgium Phone : 0032 10473004 Fax : 0032 10473471 Email : Veys-at-bota.ucl.ac.be ---------------------ooooO-----------Ooooo-------- ( ) (_)(_) ( ) ) ( ) ( (_) (_)
I«m thinking in buying a saphire knife for ultramicrotomy since they seem to be less expensive than the traditional diamond ones. I need to cut thin sections of sponges that are difficul to cut with glass knifes. Does anyone have experience with these knifes? How do they compare with diamond in terms of cutting properties and durability?
Thanks
Dr. A.P. Alves de Matos Dental Medical School Lisbon
Many of our customers are using Agfa Duoscan scanners with excellent results. These are "flatbed" type scanners but handle films in a separate drawer, similar to a negative carrier in an enlarger. The advantage of this system is not scanning through glass, eliminating the chance of Newton Rings. In addition, the Duoscan line offers high optical resolutions(up to 2500x2500ppi)and high dynamic ranges. The Umax Powerlook III is also an excellent scanner where budgets may be limited. It has 1200x2400 optical resolution and is a traditional "flatbed" design. Also new is the Linocolor 1400 with 1200x2400 resolution with a letter size scan bed.
Choosing a printer is more difficult, depending on your output needs. High end photographic printers such as the Fuji Pictrography or dyesub printers from Kodak and Sony offer top quality output but at a high price for both hardware and cost per print. For publication quality prints, these are the best. Ink jet printers continue to improve in image quality, and more importantly, long term image stability. The cost of these printers is very low although they are very slow, and still somewhat costly per print when used with the higher quality print materials. Most inkjets are also better at producing color prints than monochrome prints. Another favorite of ours is the Tektronix Phaser 850. This high quality plain paper printer uses a unique Solid Ink technology. Ink is supplied not in a liquid form but a solid blocks. Cost per print is very low and black ink is free for the life of the printer. The Phaser 850 will also handle any "office" type output such as letters and reports with the advantage of integrating images into pages instead of attaching all photos at the end.
We are currently working on a project involving blastocysts and since we aren't osmicating the samples, we also have the problem of seeing them in the resin. Embedding them in agar helps somewhat. Even though it's also relatively transparent, the larger size of the agar chunk makes it easier to see.
We perform our primary fixation, then buffer washes, then make a 2% agar solution on the hot plate. When the agar cools down enough to be quite warm to the touch (but before the gelling stage), we pipette our cells into it on a microscope slide or cover slip, then put it into the fridge to harden. It hardens almost immediately. Then we cut the piece of agar with the sample into a tiny cube and continue processing it normally.
Hope this helps.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
-----Original Message----- } From: Carrie Golash [mailto:cdg126-at-psu.edu] Sent: Thursday, June 15, 2000 9:14 PM To: Microscopy-at-sparc5.microscopy.com
Hello all -
I'm looking for advice on embedding bovine oocytes (~100 micron diameter). I'd like to embed them in araldite for probing with labelled lectins as this seems to be fairly well established in the literature. The hangup is this: we are using fairly large plastic cassettes and are having problems losing the oocytes within the volume of araldite. Does anyone know of a way around this? Is there something we can pre-embed the oocytes in to make a smaller chip that we can then embed in the larger block (that is, of course, compatible with polymerizing / clearing the araldite? Or does anyone know of an altogether different method for oocyte embedding that is more effective? Thanks in advance!
--Carrie Golash
Carrie Golash John O. Almquist Research Center Penn State University University Park, PA 16802 W: (814) 865-5896 H: (814) 692-7926 http://www.das.psu.edu/dbrc/dbrc.htm
At 11:44 AM +0100 6/16/0, A.P. Alves de Matos wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
********************* I had bought a saphire knife, years ago (early 1980's). Our lab bought it with the idea that it was a good half-way stop for a new tech who needed something better than glass. It had certain drawbacks....the edge seemed to collect debris and was more difficult to clean than a diamond, and of course it wore faster too. she used it for a while (6 months?) and then we were able to buy another diamond knife.
I haven't tried a saphire knife since, although I am partial to them in jewelry!
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
When we were working with porcine oocytes it was necessary to handle each individually so we enrobed them in agarose inside a cell of nylon net of a dark color, using a dissecting microscope. We saved enough of the excess nylon net to use as a "handle" to pick up the sample and moved it from solution to solution. This should be done after fixation, since glut fixed agarose is sometimes a problem. We also used the low temp gelling agarose, so that we had time to work. Then put it in the frig to solidify. It will then remain solid at room temp. You might get better lectin labeling using one of the acrylic resins rather than an epoxy
At 09:13 PM 06/15/2000 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Gregory W. Erdos, Ph.D. Assistant Director, Biotechnology Program PO Box 110580 University of Florida Gainesville, FL 32611
I see no reason why this type of cross sectional view cannot be achieved by sectioning with a cryostat, this is the type of use our cryostats are supplied for. If you would like more information please get back to me, I would be happy to section some samples for you to inspect.
Best Regards
Alan Bright
Bright Instrument Co.Ltd. St Margaret's Way Huntingdon PE18 6EB England
Tel No:+44 (0)1480 454528 Fax No:+44 (0)1480 456031 Email: AlanBright-at-brightinstruments.com Web Site: www.brightinstruments.com
-----Original Message----- } From: Smartech [mailto:smartech-at-javanet.com] Sent: 15 June 2000 15:03 To: To all on the list
I needed to have a cross sectional view into an adhesive layer so that I could count the layers, if possible. Since the adhesive acts like a highly viscous liquid, polishing it is out of the question. Instead, I submerged the film into liquid Nitrogen and cut it quickly w/ a scissors. The sound of the cut was more like a breaking sound. When I examined the cross sectional surface I observed a brain like surface. There were columns with a highly consistent diameter and orientation. It looked crystalline. There was zero evidence of material smearing that one would expect if one cut a material. Is the proper interpretation that these columns existed before the fracture and that the fracture occurred along the boundaries? Or is the cross sectional surface generated by a rippled distortion of an highly viscous liquid? The regularity of the surface seems to make the latter interpretation unlikely.
I would be interested in any opinions on the interpretation of the image or alternative means of cross sectioning the adhesive layer.
I could e-mail an image to anyone who is interested.
I'll ring in & say yes. I am quite happy with this combination. You get the digitized images with a much large field of view. Photo quality ink jets are cheap & in general do well. You will always find extremist in on the subjects of the infinitely best scanner & printer but here is what I bought for {9K$. AGFA Duoscan T2500 ~$4500 500MHz PC with 1/2 Gig memory & 19" hi res monitor, CD writer ~2.5K$ Epson Stylus 870 ~$300 Photo Shop, Fovea 1.0 IP software {1K$ with student ver. of PS Misc. supplies some $
For a MacPerson, my understanding is that in terms of image processing speed the Macs are 2-4x faster that PC but I don't have any benchmarks on the latest generations.
If your printing a lot, the ink jets will drain cartridges pretty quick. Down the road I will probably pick up one of the wax printers. They cost something like 3k$ but I think it is Tektronix that offers to supply all the black wax you can user for life, they are pretty fast & no secret papers are required (much cheap per BW page).
Just my thoughts before coffee.
Bruce Brinson
disclaimer... no financial interest in any companies mentioned.
tbonner 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 } -----------------------------------------------------------------------. } } Being unable to afford a digital camera for my TEM. I'm wondering if the next } best option is to get a high end scanner to scan in negatives and then print } them on a decent printer. Any advice regarding this idea and brands of } scanners and printers that are useful? Also, what image analysis systems are } user friendly? } } Dr. Thomas P. Bonner } Department of Biological Sciences } SUNY at Brockport } Brockport, NY 14420
I have been trying to replace the photocell on the Jetpolisher that I have been using. It is about 15 years old and is made by Struers. The Model is a Tenupol and the power supply is type is Polipower. Struers no longer makes replacement parts for these units. If anyone has any information concerning the photocells of this model (who I might contact to replace it or the sensitivity of the photocell) it would be greatly appreciated. Regards Kevin
} Being unable to afford a digital camera for my TEM. I'm } wondering if the next best option is to get a high end } scanner to scan in negatives and then print } them on a decent printer. ...
The next best option would be a 4x5 film scanner (~US$4k). The problem with typical film scanners is their anticipated dynamic range for photographic film, which is where TEM digital capture excels. I suggest you take a representative film and evaluate the Polaroid "4x5 Ultra". Althought I'm unfamiliar with this particular 4x5 scanner, it is the only one (I'm aware of) which is purported to scan an OD better than 3.5 (approximately 14 f/stops ... 4 f/stops per OD unit ... correct me if I'm wrong). Less expensive (~US$1.2k) would be a flat bed scanner designed for transparencies as well as hardcopy. This additional feature could be a "drawer" for film, or a optional "lid" which provides a lamp from above.
I need tips, descriptions, references regarding the preparation of cross-section specimens of the magnetic layer of magnetic recording tape for high magnfication SEM and conventional TEM observation. Thanks.
Actually the printing part has recently gotten much easier. ElectroImage (http://www.electroimage.com) is offering new technology that lets you print real grey scale images on simple inkjet printers. They have grey inks and new printer drivers.
Bill Miller
At 08:47 AM 6/16/00 -0700, George Laing wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Off hand I know I have information on the cross sectioning of hard disk media using the Tripod Polisher¨. I'm not sure if I have anything on magnetic recording tape. If you send me your mailing address, I'll send you whatever I can find that comes close.
David Henriks Vice President TEL: 800-728-2233 (toll free in the USA) South Bay Technology, Inc. +1-949-492-2600 1120 Via Callejon FAX: +1-949-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy.
Message text written by INTERNET:"EBMet-at-aol.com"-at-sparc5.microscopy.com } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
To All:
I need tips, descriptions, references regarding the preparation of cross-section specimens of the magnetic layer of magnetic recording tape for high magnfication SEM and conventional TEM observation. Thanks.
In response to Eric's question, I run a diagnostic Pathology lab at the University of South Florida, and have a one day processing schedule for kidney biopsies that calls for osmication in 1% buffered osmium at room temperature for 30 minutes. I have found that for tissue pieces in the order of 1/2 millimeter thick in one dimension the fixation is fine, and is equivalent to our routine processing osmium fixation of one hour at 4 degrees. I haven't tried this on muscle biopsies, but if they meet the thickness criteria they should be O.K. too,. Just make sure to rinse these extensively (3x 10 minutes, perhaps) in buffer to remove the excess osmium from the muscle tissue, as fluids enter and leave muscle slower because of the extensive connective tissue sheaths around the myocytes. Overosmication is a definite possibility, with subsequent tissue brittleness, if tissue is left in osmium too long, no matter what the temperature. If the tissue is too thick, uneven osmication can occur, where the outside of the tissue is well fixed and a fixation gradient is set up with poor fixation towards the center of the tissue. I observed this happening at a renal lab in Pittsburgh where I used to work. Our unstained thick sections were darker at the periphery than in the center. Another sign of this problem is lack of specimen contrast at the center of thin sections. So, Eric, as far as my experience goes, it is possible to start with 4 degree, buffered osmium, and to osmicate at room temperature for a half of an hour and get results equal to those from osmication at 4 degrees for one hour if your tissue is sufficiently thin in at least one dimension. I haven't noticed any precipitation problems with this technique, nor does the osmium discolor during fixation. Our lab has been using this technique for several years now. Take care! Ed Haller, U.S.F. Pathology
You may want to ask Struers for a users list. There may be someone out there with an older unit that is no longer being used who may be willing to give it to you for spare parts. If they can't give you a list, let me know - I think I can dig up an old list I put together of some previous Tenupol users you may be able to contact. If that doesn't work, you may want to consider upgrading to a South Bay Technology Model 550D Jet Polisher. If you have an interest in getting more information on that option, please contact me and I'll send you information.
David Henriks Vice President TEL: 800-728-2233 (toll free in the USA) South Bay Technology, Inc. +1-949-492-2600 1120 Via Callejon FAX: +1-949-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy.
Message text written by INTERNET:"kklos-at-mail.mse.ufl.edu"-at-sparc5.microscopy.com } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I have been trying to replace the photocell on the Jetpolisher that I have been using. It is about 15 years old and is made by Struers. The Model is a Tenupol and the power supply is type is Polipower. Struers no longer makes replacement parts for these units. If anyone has any information concerning the photocells of this model (who I might contact to replace it or the sensitivity of the photocell) it would be greatly appreciated. Regards Kevin {
To the person asking about the possibility of doing immunofluorescence microscopy on cells grown on plastic coverslips, someone has published a technique in BioTechniques that I saved in case I needed it. Volume24, number 6, pages 910-914, 1998 is the article titled "Mounting technique allows observation of immuno-labeled cells on plastic coverslips". The basic technique from M. F. Donohue et al involves using Thermanox coverslips on which cells are grown and immunolabeled. Following labeling, this group uses a drop of aqueous mounting medium to mount the side of the coverslip without cells on it to a glass slide. On top of this, the group then mounted a regular glass coverslip with an additional drop of aqueous mountant, and then could do their microscopy. The authors state that the inherent strong autofluorescence is greatly reduced by this technique, and the problem with the plastic not transmitting light well is overcome. Although I haven't tried the technique yet, it sounds like a simple fix for a sticky problem. I hope this is of help to you! Ed Haller, U.S.F. Pathology
Hi, We are currently in the market for a tissue processor for electron microscopy. It will mainly be used for biolgical specimens (some quite small). Although I have considerable experience with the processor sold by RMC (Ventana), I know virtually nothing about the Lynx (now being sold by EMS, I think). Any information on the advantages or disadvantages of either model (or any other one that might be out there) would be appreciated. Offline replies are welcome.
Tom Januszewski Senior Electron Microscopist Molecular and Cellular Imaging Facility UT Southwestern Medical Center at Dallas 5323 Harry Hines Blvd. Dallas, TX 75390 Email: tom.januszewski-at-email.swmed.edu
We are using the Agfa T2500 to scan TEM and SEM negatives, which also gives us the ability to scan prints. The 1200 dpi is sufficient for most needs, with 2500 dpi getting used less often and mostly with low mag images. It is currently connected to a 233 MHz Mac G3/160 Mb RAM, being replaced with a 400 MHz G4/320 Mb. The Agfa is driven by either a stand alone app. or through a plug-in that runs under most software, such as Photoshop or Object Image. You will want a lot RAM and drive space, CDR, Ord, Jaz or DVD-RAM drives. Zip drives fill far too quickly.
Photoshop is used for publication images, although some users prefer Canvas. Most of our image analysis uses the Object Image enhanced version of NIH Image. It is very easy to use. I've less experience with Image/J, but it is quickly adding capabilities and will display } 8 bits, whereas Object Image will process 16 but only displays 8 bits. Printing is either to an Epson 850, 3000, or our venerable Phaser IIsdx. BTW, Tektronix sold its printer division to Xerox.
tbonner 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 } -----------------------------------------------------------------------. } } Being unable to afford a digital camera for my TEM. I'm wondering if the next } best option is to get a high end scanner to scan in negatives and then print } them on a decent printer. Any advice regarding this idea and brands of } scanners and printers that are useful? Also, what image analysis systems are } user friendly? } } Dr. Thomas P. Bonner } Department of Biological Sciences } SUNY at Brockport } Brockport, NY 14420
--
Glen MacDonald Virginia Merrill Bloedel Hearing Research Center Box 357923 University of Washington Seattle, WA 98195-7923 glenmac-at-u.washington.edu (206) 616-4156 (206) 616-1828 fax *********************************************************** C:} The box said "Requires Windows95 or better". So I bought a Macintosh. ***********************************************************
} I«m thinking in buying a saphire knife for ultramicrotomy since they seem to } be less expensive than the traditional diamond ones. I need to cut thin } sections of sponges that are difficul to cut with glass knifes. Does anyone } have experience with these knifes? How do they compare with diamond in terms } of cutting properties and durability? } } Dr. A.P. Alves de Matos } Dental Medical School } Lisbon
Unfortunately, you get what you pay for. Sapphire does NOT have the durability of diamond and will be damaged by the sponge spicules. And, to my knowldge, they can't be resharpened.
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
It could be a solution if you mainly want an alternative to silver prints, or to acquire images for Powerpoint presentations. However, many of the really useful features of a digital camera on a TEM are unavailable if you adopt this approach, namely instant verification of image capture, greyscale expansion, image averaging, online analysis, and many more. Also, you still need to allocate some space to a darkroom.
I looked at large-format transparency scanners a couple of years ago, when there was little of this kind in the market, and concluded that their combinations of bit depth, pixels per inch and sensitivity and dynamic range at the high-density end of the negative (i.e. highlight detail) was close to what was required if the objective was merely to obtain publication quality images from a large proportion of negative area (these images require optimised contrast and crispness, but being small do not demand much resolution), but really inadequate for scanning of image details (organelles, molecules), for dense exposures and highlights, and for high contrast subjects like replicas. Most of these scanners appeared to be optimised for scanning positive images (large format colour transparencies) where discrimination of detail in the extreme shadows is not top priority. However, this becomes a major shortcoming when dealing with negatives.
I would certainly like to know whether anyone feels that there is an adequate solution available today.
I don't think user friendliness is the most useful criterion for discriminating between image analysis packages. This is in any case a fairly subjective property, depending very considerably on the computer - literacy of the user. Most IA packages (analySIS, Optimas, etc) are GUI-based systems, and therefore are reasonably intuitive. One of the features that differentiates them is the balance between the provision of off-the-peg analysis solutions and programmability. The range of tasks demanded of an IA package is potentially so great that there is little alternative but to evaluate them and see if they suit your needs. However, I warn you that your needs are likely to evolve. What seems like a simple and user- friendly solution today will probably feel like a very limited and inflexible one tomorrow if it has insufficient functionality and programmability, and these things unavoidably add complexity.
Chris Jeffree
Date sent: Thu, 15 Jun 2000 21:14:18 -0500 To: Microscopy-at-sparc5.microscopy.com } From: tbonner {tbonner-at-brockport.edu}
tbonner said: } Being unable to afford a digital camera for my TEM. I'm wondering } if the next best option is to get a high end scanner to scan in } negatives and then print them on a decent printer.
Using a flat bed scanner over a digital camera for TEM images is definately your best option budget wise. We use an Agfa Duoscan scanner with great success. it is very versitile, we use it to scan gels, scan TEM negatives and scan old prints. It can scan slides for powerpoint quality presentations also. So check with Agfa to see their latest lineup.
For printers inkjets work great when combined with photoquality papers. Any (Epson HP & Cannon) 300dpi or higher color ink jet will give decent images suitable for posters. For publications dye sublimation printers work well but get one that uses a cartridge (one piece) to replace empty media. For proofing look for laser printers that are at least 1200dpi with extra ram (64meg on the printer is a nice number to start with) and large toner cartridges, graphic images burn a lot of toner. Look for a printer that will print alot before replacement of the toner.
For software, Photoshop is a good choice. Before you spend alot on a venders image analysis package try some of the freeware out there like NIH image and Image J both from the NIH website. If you can't get the free stuff to work, then spend the extra money. We use Image J here and it works well.
For computers (Apple or PC) consider one scanner with a SCSI interface with need a SCSI card (avoid parallel port and universal serial bus (USB) scanners unless you like coffee breaks). So you need one computer to hook up to the scanner but Ideally you would also have three printers (inkjet, Laser & Dye sub or comparable) But Inkjets are cheap make your users get one and maintain it. I bet somewhere in your department is a networked laser printer so print to a networked printer elsewhere. Keep the high end printer (Dye sub/thermal printer close by) Invest in removable media that others can use like a CD-r (HP or Plextor) and a zip drive at the bare minimum.
Hope this helps Jon Ekman Associate Research Specialist Deptartment of Biological Sciences University of Wisconsin-Milwaukee phone W:414.229.6471 Web1 http://www.graffitimasters.com Web2 http://www.uwm.edu/~jekman
Embed it, Elliot and diamond knife section it in an ultramicrotome, using the block face for the FE-SEM examination and the ultrathin sections (20-200 nm thick) for examination in the TEM. A good reference is Ho et al, Specimen Preparation for TEM of Materials, Mat. Res. Soc. Symp. Proc., vol. 115, pp. 149-154 (MRS, Pittsburgh, 1988). You may have to search around for someone who can do this for you, but it is far and away the best technique for your needs.
Tom Malis Group Leader - Characterization Materials Technology Laboratory Natural Resources Canada (Govt. of Canada) 568 Booth St., Ottawa, Canada ph. 613-992-2310 FAX 613-992-8735 email: malis-at-nrcan.gc.ca
---------- From: "EBMet-at-aol.com"-at-sparc5.microscopy.com [SMTP:"EBMet-at-aol.com"-at-sparc5.microscopy.com] Sent: Friday, June 16, 2000 11:35 AM To: microscopy-at-sparc5.microscopy.com Subject: re: Cross-section Preparation of Magnetic Recording Tape
------------------------------------------------------------------------ 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 need tips, descriptions, references regarding the preparation of cross-section specimens of the magnetic layer of magnetic recording tape for high magnfication SEM and conventional TEM observation. Thanks.
I am ion milling gold. nearly 100 nm. I want to know how long it takes to make it thin enough to be transparent under TEM. Thanks.
Gen ****************************************************************** Gen Pei Department of Materials Science and Engineering Cornell University 328 Thurston Hall tele: (607)255-5177 fax:(607)255-2365 gp35-at-cornell.edu
a) So far as I know they are not made any more and have not been made for at least ten years, and
b) The economics have changed drastically from twenty years ago when the sapphire knife did enjoy a bit of popularity. In real terms, diamond knives, now because of the competition from Microstar have drop significantly from what they once were, perhaps 50%, so whatever pricing advantage there was at one time, did not exist any more. So the Japanese company that made them discontinued their production. It was called "Saphatome" or something like that. Ted Pella would probably know their history, perhaps better than I do.
Also, because of your interest in education, take a look at www.microscopy-advantage.com . Tell me what you think. Attendees at the coming meetings of APEM, EUREM and MSA will automatically receive a CD in their registration materials. If you would like a copy, send me your UPS address and I will make sure that one gets sent to you. But it will "work" exactly as if you were on line. --- Original Message --- Caroline Schooley {schooley-at-mcn.org} Wrote on Fri, 16 Jun 2000 11:47:54 -0700 ------------------ ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} I´m thinking in buying a saphire knife for ultramicrotomy since they seem to } be less expensive than the traditional diamond ones. I need to cut thin } sections of sponges that are difficul to cut with glass knifes. Does anyone } have experience with these knifes? How do they compare with diamond in terms } of cutting properties and durability? } } Dr. A.P. Alves de Matos } Dental Medical School } Lisbon
Unfortunately, you get what you pay for. Sapphire does NOT have the durability of diamond and will be damaged by the sponge spicules. And, to my knowldge, they can't be resharpened.
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
----- Sent using MailStart.com ( http://MailStart.Com/welcome.html ) The FREE way to access your mailbox via any web browser, anywhere!
} It could be a solution if you mainly want an alternative to silver prints, } or to acquire images for Powerpoint presentations. However, many } of the really useful features of a digital camera on a TEM are } unavailable if you adopt this approach, namely instant verification of } image capture, greyscale expansion, image averaging, online } analysis, and many more. Also, you still need to allocate some } space to a darkroom.
Why not just outsource the processing of the film? Depending on where one resides/operates, there are typically numerous professional and non-professional labs which will do same day development of b/w film. I do this for 4x5 cut sheet film and 120/220 roll film from a regular camera and from the SEM recording camera. Unless there is some overriding need or requirement for an on-site darkroom, why not just send the film out whenever it is needed? I could see the rationale for an on-site facility if the TEM was producing hundreds of negs per day or perhaps per week. Then it is a make-buy decision regarding in-house or out-house processing.
If one is concerned about whether a shot will turn out (instant verification), just shoot a couple more sheets or frames bracketed around the "optimum/normal" exposure time. The cost of the film and processing is way too low to justify a high cost digicam for TEM. SEM imaging is of course a totally different matter.
I find that grey scale expansion is not the sole domain of the digicam. In a neg, additional information is there--but typically the eye cannot see it. This is where image analysis and image processing programs are very beneficial.
} I looked at large-format transparency scanners a couple of years } ago, when there was little of this kind in the market, and concluded } that their combinations of bit depth, pixels per inch and sensitivity } and dynamic range at the high-density end of the negative (i.e. } highlight detail) was close to what was required if the objective was } merely to obtain publication quality images from a large proportion } of negative area (these images require optimised contrast and } crispness, but being small do not demand much resolution), but } really inadequate for scanning of image details (organelles, } molecules), for dense exposures and highlights, and for high } contrast subjects like replicas. Most of these scanners appeared to } be optimised for scanning positive images (large format colour } transparencies) where discrimination of detail in the extreme } shadows is not top priority. However, this becomes a major } shortcoming when dealing with negatives.
Discrimination of detail in shadows is a major concern for users of transparencies. This is why they seek high D rated scanners. I typically scan negative and transparencies as transmitted RGB or greyscale. This is because I find that the scanner picks up more detail across the whole image when scanned as a tranny.
} [snip]
} and see if they suit your needs. However, I warn you that your } needs are likely to evolve. What seems like a simple and user- } friendly solution today will probably feel like a very limited and } inflexible one tomorrow if it has insufficient functionality and } programmability, and these things unavoidably add complexity. } } Chris Jeffree
I agree that needs may evolve. That means that when making the initial purchase of an image analysis program, it should be flexible enough to allow custom augmentation. Most of the higher end ones do. But it may turn out that one program alone is not as good as two different programs--each being good at different aspects of image analysis.
gary g.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Modern surfers use PC boards. You can too at http://photoweb.net ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Tom Malis wrote: ====================================================== Embed it, Elliot and diamond knife section it in an ultramicrotome, using the block face for the FE-SEM examination and the ultrathin sections (20-200 nm thick) for examination in the TEM. A good reference is Ho et al, Specimen Preparation for TEM of Materials, Mat. Res. Soc. Symp. Proc., vol. 115, pp. 149-154 (MRS, Pittsburgh, 1988). You may have to search around for someone who can do this for you, but it is far and away the best technique for your needs. ======================================================
This has been our experience too, however we add the following to the preparation protocols:
a) We coat one side of the recording tape with, say, Pt, the other side with Al, because once in the TEM, it is important to validate that i] nothing has fractured off during the ultramicrotomy and ii] you can keep straight which side is which for the asymmetric cross-section. If the two metallization lines are present in the TEM, with embedding resin on the other side, you can be certain you are seeing the entire cross-section. If one is missing, you might not have the entire cross-section.
b) For looking at the "faced-off-piece", we suggest an ever so slight amount of oxygen plasma etching, in order to bring out a bit more contrast between the ferrite or other inorganics from the matrix polymer. The inorganics stand up like little "mesas" in the desert, giving greatly enhanced contrast. Since you now have an element of three dimensional nature to the same, you can gain some insight into orientation, something that would not otherwise be possible
Disclaimer: If you are looking for someone to do this kind of work, look no further, we have been doing this kind of sample preparation for clients on a contract basis since the early 1970's! Our own Plasma Prep II plasma etcher would do the described etching on the faced-off-piece after about 120 seconds of exposure.
Chuck
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And if you're satisfied with prints at 720x1440 dpi, Epson has just made a major leap in ink and paper longevity; read about it at http://www.epson.com/whatsnew/ygtsi/lightfast.html http://www.wilhelm-research.com/ . Unfortunately, the new ink cartridge won't fit old (as in last year's) printers. Oh well - what else might I spend $370 on?
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
List Recipients: I am posting this message at the request of Joe Lester and all correspondence should be sent to him. Dave Audette david.audette-at-sylvania.com
JOB OPENING Scanning Electron Microscopist in Analytical Laboratory ( Beverly, MA)
OSRAM Sylvania, Inc. 71 Cherry Hill Drive Beverly MA 01915
DESCRIPTION: Structural and elemental characterization of materials used in incandescent, fluorescent and discharge lamps, especially by optical and electron microscopy. Failure analyses of lamps and lighting components. Technical problem solving as a member of a team. Oral and written communication of results and conclusions with client population.
POSITION REQUIREMENTS: Competence in optical and electron microscopy of materials including EDS. An understanding of failure analysis. Ability to work independently and/or in a team and to communicate effectively.
EDUCATION AND EXPERIENCE REQUIREMENTS: B.S., or higher, in Materials Science, Chemistry, or Physics. 2-5 years experience in SEM/EDS. Experience with lamp components is desirable.
Please send a resume to
Dr. Joe Lester Technical Assistance Lab OSRAM Sylvania Inc. 71 Cherry Hill Drive Beverly, MA 01915 e-mail: joe.lester-at-sylvania.com
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G'day All, In my experience Sapphire Knives are an excellent replacement for glass knives when working with soft materials. Sapphires are much softer than diamond and are more easily damaged than diamond. Sponges are NOT soft tissue, they contain very hard inorganic salt spicules that damage both glass and sapphire knives. Regards JVN
Leona Cohen-Gould 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 } -----------------------------------------------------------------------. } } At 11:44 AM +0100 6/16/0, A.P. Alves de Matos 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 All } } } } } } I«m thinking in buying a saphire knife for ultramicrotomy since they seem to } } be less expensive than the traditional diamond ones. I need to cut thin } } sections of sponges that are difficul to cut with glass knifes. Does anyone } } have experience with these knifes? How do they compare with diamond in terms } } of cutting properties and durability? } } } } } } Thanks } } } } Dr. A.P. Alves de Matos } } Dental Medical School } } Lisbon } } ********************* } I had bought a saphire knife, years ago (early 1980's). Our lab bought it } with the idea that it was a good half-way stop for a new tech who needed } something better than glass. It had certain drawbacks....the edge seemed } to collect debris and was more difficult to clean than a diamond, and of } course it wore faster too. she used it for a while (6 months?) and then } we were able to buy another diamond knife. } } I haven't tried a saphire knife since, although I am partial to them in } jewelry! } } 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
-- **************************************************** John V Nailon Operations Manager Centre for Microscopy and Microanalysis The University of Queensland St. Lucia Queensland 4072 Phone: +61-7-3365-4214 Fax: +61-7-3365-4422 WWW: http://www.uq.edu.au/nanoworld/allstaff.html#Nailon ****************************************************
I had a argument with my husband, he says he is going to the Kunming for an EM meeting,
http://www.iphy.ac.cn/microsc/IKSM.html
He says there are many good scientists going which I believe with doubt. But I think he is going for a sight seeing. There is a Chinese saying: The mountains and waters in Guilin are the most beautiful ones under the sky. I have asked him to bring me, he agreed but unable to get the same airline ticket (UA fully booked). Any body is going and knows alternative airlines, please contact me. Thanks a lot.
I have two DVD-RAM drives and 15 media (Type I and II). I have found that these are riddled with write & read errors. Be careful when using this storage media.
My main unit is a Panasonic LF-D101 (SCSI) and the second one is the same. The third is a Matshushita ID unit.
Scandisk will report either many errors that are fixed or no errors. Either way, the media/drive will write faulty file contents.
Hi Hao, We have in stock (100), (110) and (111) MgAl2O4 substrates with 7 angstrom finish ready for laser ablation or other kinds of epitaxial film deposition. Contact me for info regarding perovskite lattice matching. Best regards, Mike Urbanik www.crystalguru.com
{ { Subj: information about MgAl2O4 Date: 6/15/00 3:33:44 PM Eastern Daylight Time From: haoli-at-glue.umd.edu (Hao Li) To: Microscopy-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 -----------------------------------------------------------------------.
Hi, All:
This one is not really related to microscopy. But since I am a TEM guy, I hope I can get some help here.
Does anybody have some information about MgAl2O4 as a substrate for perovskite films? I know it is not often used, so I am worndering if it has a major disadvantage so that nobody is using it. Any references would be welcome.
Hi All, I processed a series of pellets of yeast for a client using a glut-pfa fix, osmium, dehydration through ethanols, and a day & a half step-wise infiltration into Spurr's resin (1:1 with Ethanol for 4 hr, pure resin overnight under light vacuum, then fresh for 3 more hours, then embed in fresh). Polymerization was overnight at 60C. About half the blocks were soft and had to be returned to the oven for a prolonged polym. (over the weekend). I was able to get sections from each of the 10 samples, but in the 'scope, many of these looked a bit like swiss cheese. those that were not lacy exhibited areas where the resin pulled away from the cell coats of the yeasts. Clearly something went wrong with the infiltration/polymerization. I used the same batch of resin for other things, and its fine.
Does anyone out there have experience with yeast? Any suggestions? I'd like to give this peson some usable data!
Thanks in advance, 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
My only thought is that your infiltration/dehydration were too short, or your 100% ethanol had absorbed too much water. My understanding is that Spurr's is very sensitive to small quantities of water, with soft blocks and holes being symptoms of incomplete dehydration. Sometimes we extend the dehydration through 3 changes of 100% ETOH with molecular sieves, then on through 2-3 changes of propylene oxide. Infiltration is usually 1:2 PO:Resin, followed by 1:1, 2:1, then a couple changes of pure resin overnight or for 4-8 hours, then final embedding in another change of pure resin.
I don't know if yeast is more problematic than other things in this respect, however.
Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
-----Original Message----- } From: Leona Cohen-Gould [mailto:lcgould-at-mail.med.cornell.edu] Sent: Monday, June 19, 2000 8:47 AM To: Microscopy-at-sparc5.microscopy.com
Hi All, I processed a series of pellets of yeast for a client using a glut-pfa fix, osmium, dehydration through ethanols, and a day & a half step-wise infiltration into Spurr's resin (1:1 with Ethanol for 4 hr, pure resin overnight under light vacuum, then fresh for 3 more hours, then embed in fresh). Polymerization was overnight at 60C. About half the blocks were soft and had to be returned to the oven for a prolonged polym. (over the weekend). I was able to get sections from each of the 10 samples, but in the 'scope, many of these looked a bit like swiss cheese. those that were not lacy exhibited areas where the resin pulled away from the cell coats of the yeasts. Clearly something went wrong with the infiltration/polymerization. I used the same batch of resin for other things, and its fine.
Does anyone out there have experience with yeast? Any suggestions? I'd like to give this peson some usable data!
Thanks in advance, 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
Dear Gen Pei, I have looked at gold contacts, lifted from an electronic device, that were supposed to be 100 nm thick. I could see the structure clearly at 200 kV. At 10:57 PM 6/16/00 -0400, you wrote:
} HI, } } I am ion milling gold. nearly 100 nm. I want to know how long it } takes to make it thin enough to be transparent under TEM. Thanks. } } Gen
Regards, Mary
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-interchg.ubc.ca
I was just visited by one of our EH&S folks who wanted to know why I had 1,2 dichloroethane.
Seems they track purchases and I bought some last year to make formvar films.
1,2 dichloroethane is on their bad list as a carcinogen. He was actually here to figure out how much might be going up the fume hood so he could make a report to the local air quality agency. But as we talked, it seemed like it would be better to not have the stuff around.
Anyone have experience with formvar in chloroform? I read it works but have never tried it. According to our EH&S guys, chloroform would be better than 1,2, dichloroethane.
BTW we are in California and must abide by some pretty strict rules, it may seem like they are going overboard, but they are just trying to do their job.
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
{P} Good day to all on the listserver, {/P} {P} {/P} {P} I have a person in my department who is interested in processing horse sperm for TEM. Does anyone who does this routinely be willing to give me some pointers as to how to process them? I've only worked with muscle and brain tissue so this is kinda new - I have processed cells from cell culture for TEM would it be the same procedure? {/P} {P} {/P} {P} Thanks so much, {/P} {P} Connie Cummings, DVM {/P} {P} Instructor Anatomic Pathology {/P} {P} Department VBP {/P} {P} Oklahoma State University {/P} {P} {/P}
Hi Y'all: We are looking for a for-hire independent FIB company that has experience in preparing TEM cross-sections of semiconductors. Please contact me if you do this, or know of a lab that does. Regards, Michael Coviello Lab Manager Materials Science University of Texas at Arlington
Thanks to everyone for your helpful comments. I'm taking another stab at it with smaller pellets, longer times and the addition of prop. ox. steps after the ethanol. I had pretty much decided to do all that anyway, but its nice to gets confirmation ofone's ideas!
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
Try FIBICS in Ottawa, Canada. Contacts are Mike Phaneuf or Louise Weaver. Their telephone number is 613-860-0861. email: mphaneuf-at-fibics.com or lweaver-at-fibics.com
-----Original Message----- } From: Mike Coviello [mailto:coviello-at-mae.uta.edu] Sent: Monday, June 19, 2000 4:12 PM To: listserver
Hi Y'all: We are looking for a for-hire independent FIB company that has experience in preparing TEM cross-sections of semiconductors. Please contact me if you do this, or know of a lab that does. Regards, Michael Coviello Lab Manager Materials Science University of Texas at Arlington
Hi Lee, We follow a very similar protocol to yours with the exception that we use propylene oxide rather than ethanol for infiltration. PO : Spurrs 1:1 1 hour PO : Spurrs 1:3 1- 2 hours Spurrs 1 - 2 hours Spurrs overnight (we do not infiltrate under vacuum) fresh Spurrs 1 - 2 hours polymerization at 60C 48 hours
We had the problem you describe when we tried to embed yeast in EPON equivalents. Switching to Spurrs was the fix for us. Frank
At 09:46 AM 6/19/00 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi Leona, The standard way for dealing with yeast cells is to remove the cell wall. This is done with glusulase (fancy name for snail guts) and or lyticase (both from Sigma). If the wall cannot be removed for experimental reasons (i.e. study of the plasma membrane/cell wall interface) then you need to modify the carbohydrate linkages of the cell wall to make the wall more permeable. This can be done by treating the cells, after fixation, with 1% sodium metaperiodate for about 15 minutes. However, most researchers simply wishing to examine yeast morphology remove the wall because this not only improves inflitration of the resin it also allows more extraction of the cytoplasm and thus makes it easier to resolve structures and membranes within the ribosome rich yeast cell. A classic protocol, by Byers and Goetsch, can be found in Vol 194, Methods in Enzymology (AKA Guthrie and Fink) pg 602. I strongly encourage you to read this article as yeast can be very problematic. You also want to use the Hard Spurrs formulation and use 100% acetone (or propylene oxide) as the last dehydration step before going into 1:1 resin. I have not observed any difference in the ultrastructure of cells embedded in Spurrs vs. polybed 812 (when the wall is removed). If you want to do immuno-EM you might find our protocol useful; checkout: http://genome-www.stanford.edu/group/botlab/protocols/EM_protocol.pdf.
Jon Mulholland Genetics Dept Stanford University School of Medicine Stanford, CA 94305-5120
On Mon, 19 Jun 2000, Leona Cohen-Gould 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 All, } I processed a series of pellets of yeast for a client using a glut-pfa fix, } osmium, dehydration through ethanols, and a day & a half step-wise } infiltration into Spurr's resin (1:1 with Ethanol for 4 hr, pure resin } overnight under light vacuum, then fresh for 3 more hours, then embed in } fresh). Polymerization was overnight at 60C. About half the blocks were } soft and had to be returned to the oven for a prolonged polym. (over the } weekend). I was able to get sections from each of the 10 samples, but in } the 'scope, many of these looked a bit like swiss cheese. those that were } not lacy exhibited areas where the resin pulled away from the cell coats of } the yeasts. Clearly something went wrong with the } infiltration/polymerization. I used the same batch of resin for other } things, and its fine. } } Does anyone out there have experience with yeast? Any suggestions? } I'd like to give this peson some usable data! } } Thanks in advance, } 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 } } }
Formvar in chloroform works well. I don't think we have ever tried dochloroethane.
I have had to do the job myself recently and made a minor discovery - the film seems to stick very well to acid washed slides! So, not so cleverly clean. Then the second trick - float the film soon after it has dried, within a minute. Then it seems to work better.
Keith
_______________________ Keith Ryan (Dr) Marine Biological Association Citadel Hill Plymouth Devon PL1 2PB England
Dear all, Thanks to all who responded to my enquiry and gave me useful advice concerning the preparation of cross-sections of ceramic thin films.
Hopefully, I should now be able to prepare some better thin film specimens using one or more of the suggestions that I received.
Thanks again
Best wishes
===== Ian MacLaren Beijing Laboratory of Electron Microscopy Chinese Academy of Sciences, P.O. Box 2724 100080 Beijing China General Email: ian.maclaren-at-physics.org Work (esp. large attachments): maclaren-at-image.blem.ac.cn
____________________________________________________________ Do You Yahoo!? Get your free -at-yahoo.co.uk address at http://mail.yahoo.co.uk or your free -at-yahoo.ie address at http://mail.yahoo.ie
12th Scandinavian Conference on Image Analysis SCIA 2001
June 11-14, 2001 in Bergen, Norway
Sponsored by: IAPR (The International Association for Pattern Recognition) http://www.iapr.org
First Announcement and Call for Papers:
http://www.ux.his.no/scia2001/
Invitation to the 12th SCIA. Following the previous conferences in Greenland, SCIA 2001 - the 12th Scandinavian Conference on Image Analysis - will be held in Bergen on the west coast of Norway. The conference is arranged by the Norwegian Society for Image Processing and Pattern Recognition (NOBIM) and sponsored by the International Association for Pattern Recognition (IAPR). The conference venue is Grieghallen, located in the city centre.
Scientific Program: The conference will offer internationally acclaimed speakers in plenary talks and parallel sessions with selected oral presentations and posters. The conference language is English. The different presentations will cover unpublished theoretical or applied research results.
Invited Speakers: Professor Theo Pavilidis, State University of New York at Stony Brook, USA: "History of Image Analysis" Professor Josef Begün, University of Halmstad, Sweden: "Biometric Person Authentication" Professor Matti Pietikäinen, University of Oulu, Finland: "Machine Vision and Media Processing" Associate Professor Torbjřrn Eltoft, University of Tromsř, Norway: "Neural Network approaches to Cluster-Detection-and-Labelling"
In addition, we are working to find an invited speaker for the subject: "Images in the future mobile terminals".
Pre-conference Workshop/Tutorial: A set of pre-conference half-day workshops/tutorials will be held on June 11, 2001: 1. ICA (Independent Component Analysis): Professor Erkki Oja, Helsinki University of Technology, Finland. 2. Data fusion: Professor Jon Atli Benediktsson, University of Iceland.
Paper submission and registration for presenting authors: Only full papers in English will be accepted, and the length should not exceed eight pages. All papers will be refereed by two reviewers for publication in the conference proceedings. Please send four copies of your paper to:
SCIA2001, Department of Electrical and Computer Engineering, Stavanger University College, P.O.Box 2557 Ullandhaug, N-4091 Stavanger, Norway
Important dates:
Paper submission deadline: November 6, 2000 Notification of acceptance: January 19, 2001 *Camera-ready copy: March 19, 2001
*Camera-ready copy must be accompanied by registration and payment by presenting author. The cover page must contain: * Title of the paper * Name(s), complete address and e-mail for the author(s) * Brief abstract (150-200 words) * Keywords describing the main subject of the paper (3-5 words)
* Author's opinion on whether the paper is most suitable for oral or poster presentation * Name and address for correspondence
Papers considerably longer than the final size, risk being rejected. The fee for one or two extra pages is NOK 500 per page. The fee for colour illustrations is NOK 4000 per page. The decision on oral or poster presentation will be taken solely on suitability, not on paper quality. Paper submission information is available at http://www.ux.his.no/scia2001/, where the LaTeX style file and an example file in the recommended two-column LaTeX format is available.
Enquires: If you have scientific questions, please contact Ivar.Austvoll-at-tn.his.no. The program with further information about registration and payment will be send to you February 1, 2001. If you occasionally have seen this announcement, and want to have the program sent to you, please contact scia-at-plus-convention.no.
Program Committee: Dr. Ivar Austvoll (Chairman), Norway Prof. Jussi Parkkinen, Finland Prof. Fritz Albregtsen, Norway Dr. Alfred Hanssen, Norway Prof. Gunilla Borgefors, Sweden Dr. Anne Solberg, Norway Dr. Bjarne Ersbřll, Danmark
Organized by: Norsk forening for bildebehandling og mřnstergjenkjenning (NOBIM) (Norwegian Society for Image Processing and Pattern Recognition) http://www.nobim.no/
SCIA2001 web site:
http://www.his.no/scia2001/ or http://www.ux.his.no/scia2001/
At 12:42 PM -0500 6/19/0, Connie A Cummings wrote:
} Good day to all on the listserver, } } I have a person in my department who is interested in processing horse } sperm for TEM. Does anyone who does this routinely be willing to give me } some pointers as to how to process them? I've only worked with muscle and } brain tissue so this is kinda new - I have processed cells from cell } culture for TEM would it be the same procedure? } } Thanks so much, } Connie Cummings, DVM } Instructor Anatomic Pathology } Department VBP } Oklahoma State University **************************** Connie, Assuming that your colleague will bring you a semen sample, and that orientation is not critical, you can spin the sperm to a pellet and treat it like any other cell pellet. I've done various rodent, marsupial and human sperm samples this way. Once at the microscope, you will have to hunt around a bit to fine the appropriate views (head, mid-piece, tails,etc), but since there are so many cells in the pellet, I've always found what we wre looking for. Its the easiest way to go. Good luck, 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
I am posting this question to see if we can get some help for our building engineer to deal with problems we are having with our SEM (JEOL T330A).
We suspect that we are having a problem with the building power supply to our SEM. Other folks in our building using other types of equipment have found it necessary to use power conditioners or UPS systems to run their equipment - these systems seem to do the trick for them.
We are trying this approach with the power supply to our SEM, but find that there are some problems. When using the UPS system, we have to connect it up, put it on bypass, start up the SEM, then switch the UPS over after that - a real nuisance. If we don't do this, the SEM won't start.
Does anyone else have this problem, and if so, has found an easy solution? Please contact me offline if you have any suggestions and other words of wisdom. Thanks in advance. Also, thanks for all the help you as a group have given me when I posted previous questions - sometimes I forget to say thank you.
Paula.
Paula Allan-Wojtas Research Scientist - Food Microstructure Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
This is a preliminary message to all Gatan users who will be attending the Microscopy and Microanalysis Meeting in Philadelphia this August (13th to the 17th). I think it has become time to form a User's Group to discuss the level of service and support that we are receiving from Gatan. We should determine where the company should be focusing its efforts and lobby them to correct problems that are most important to us. Please let me know if you are interested in attending such a meeting so I can gauge whether it should be held, and how large a conference room I would need to reserve in Philly.
Note, Gatan representatives are encouraged to attend this meeting, but it will be a user meeting run by the users.
I will not rant and rave here in a completely open forum, as I believe it would be unfair. If you have concerns or comments on this subject, whether or not you are attending M&M2000, please contact me directly. Do NOT reply to the list, check the "To:" header before sending your message, it should say "jfmjfm-at-engin.umich.edu" only.
Thank You.
John Mansfield.
Disclaimer: Opinions expressed in this message are my own personal ones and do not represent necessarily those of my employers.
--
Dr. John Mansfield CPhys MInstP North Campus Electron Microbeam Analysis Laboratory 417 SRB, University of Michigan 2455 Hayward, Ann Arbor MI 48109-2143 Phone: (734) 936-3352 FAX (734) 763-2282 Cellular Phone: (734) 358-7555 (Leaving a phone message at 936-3352 is preferable to 358-7555) Email: jfmjfm-at-engin.umich.edu URL: http://emalwww.engin.umich.edu/people/jfmjfm/jfmjfm.html Location: Lat. 42ˇ 16' 48" Long. 83ˇ 43' 48"
Does anyone have experience taking specimens from acetone into LR Gold resin? Some of the brochures on LR White seem to recommend against it, but we are hoping it may be OK for LR Gold. We are doing freeze substitution through acetone for EM-immunocytochemistry.
Thanks in advance for your help. David H. Hall Center for C. elegans Anatomy Department of Neuroscience 1410 Pelham Parkway Albert Einstein College of Medicine Bronx, NY 10461
Sounds like the UPS capacity (VA) is sufficient to run the system, but under rated for the start-up surge. One solution would be a higher capacity UPS. One sized to handle the surge load, however, could be quite large and expensive.
If the nature of your problems is related to line noise, but not voltage levels, you might consider an "ultra isolation transformer" and experiment with various grounding options to minimize interference.
If voltage flucuations are the problem, investigate a "ferro resonant" transformer to stabilize the line. This transformer *should* be somewhat more economical than a similar capacity UPS. BEWARE: These devices produce a loud hum. You don't want it in the same room without some sort of noise attenuation.
.Haven't checked relative pricing, but here is a typical link: http://www.sola-hevi-duty.com/
Another posibility... Can you seperate the vacuum pump(s) supply from the electronics, powering the pumps directly and using your UPS for only the electronics? That may lessen the start-up load enough for the UPS to funcion normally.
Woody White McDermott Technology, Inc.
The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi, all,
I am posting this question to see if we can get some help for our building engineer to deal with problems we are having with our SEM (JEOL T330A).
We suspect that we are having a problem with the building power supply to our SEM. Other folks in our building using other types of equipment have found it necessary to use power conditioners or UPS systems to run their equipment - these systems seem to do the trick for them.
We are trying this approach with the power supply to our SEM, but find that there are some problems. When using the UPS system, we have to connect it up, put it on bypass, start up the SEM, then switch the UPS over after that - a real nuisance. If we don't do this, the SEM won't start.
Does anyone else have this problem, and if so, has found an easy solution? Please contact me offline if you have any suggestions and other words of wisdom. Thanks in advance. Also, thanks for all the help you as a group have given me when I posted previous questions - sometimes I forget to say thank you.
Paula.
Paula Allan-Wojtas Research Scientist - Food Microstructure Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
HI: For the second year in a row I am having to have our thin window (low element type-brand name with held) replaced. We have the EDS connected to a TOPCON SM-500 environmental SEM. I look at a lot industrial dusts for particle size and composition and the manuf. of the EDS system says they poke holes in the polymer window. Has anyone else have this problem? I have elected to have a thin beryllium installed this time since my boss is upset about spending $7000 every year plus the downtime and since low element detection is not critical . Thanks Terry Ellis Hallmark Cards Inc.
Does anyone have suggestions for encapsulating cell pellets in agarose?
I am working with flatfish cells but the pellets don't look cohesive enough to withstand washing, dehydration etc. We have some Type I agarose (Sigma, gel temp 36C, melting temp. 86C). I plan to post-fix in 1% osmium tetroxide followed by 1% aqueous uranyl acetate then embed in Spurr's resin.
As this is my first time working with cells, any help will be greatly appreciated.
Thank you very much, Carla Aiwohi Western Fisheries Research Center Seattle, WA
The particulate (have also heard) can "shoot" holes in a thin window. My former SEM/EDS was an Etec with a LARGE roughing system and a turret detector with Be, UTW, and "open" positions. Both windows lasted the 15 odd years it was in use before replacement. An important point is to evacuate and vent slowly so as to not accelerate the particles into the window.
I added a manual valve to the Etec between the roughing system and the chamber. With the automatic valves closed, I could slowly open the manual valve to rough the chamber down to a point where any particulate is not disturbed, close the valve, then switch to automatic to finish the evacuation. Venting gas was from my cryo of liquid nitrogen (pretty dry!) which I pressure/flow controlled for similar results on venting.
My modification was driven by the need to avoid damage to fluffy ceramic specimens, but worked well to protect the detector also.
Have not added such a feature to the new SEM/Thin Window EDS system, but (for now) the chamber is pristine and the fluffy specimens are not part of the work mix.
Woody White McDermott Technology, Inc
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HI: For the second year in a row I am having to have our thin window (low element type-brand name with held) replaced. We have the EDS connected to a TOPCON SM-500 environmental SEM. I look at a lot industrial dusts for particle size and composition and the manuf. of the EDS system says they poke holes in the polymer window. Has anyone else have this problem? I have elected to have a thin beryllium installed this time since my boss is upset about spending $7000 every year plus the downtime and since low element detection is not critical . Thanks Terry Ellis Hallmark Cards Inc.
My guess is that the SEM inititally draws power that exceeds the ratings of the UPS. It is probably due to the rotary pumps that can take up to ten amps upon start up.
The alternatives are to increase the power rating of the UPS or rewire the rotary pump so it draws it's power from the line and not through the UPS (the pumps is not that sensitive anyway).
Good Luck,
Earl weltmer
Paula Allan-Wojtas 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, all, } } I am posting this question to see if we can get some help for our building engineer to deal with problems we are having with our SEM (JEOL T330A). } } We suspect that we are having a problem with the building power supply to our SEM. Other folks in our building using other types of equipment have found it necessary to use power conditioners or UPS systems to run their equipment - these systems seem to do the trick for them. } } We are trying this approach with the power supply to our SEM, but find that there are some problems. When using the UPS system, we have to connect it up, put it on bypass, start up the SEM, then switch the UPS over after that - a real nuisance. If we don't do this, the SEM won't start. } } Does anyone else have this problem, and if so, has found an easy solution? Please contact me offline if you have any suggestions and other words of wisdom. Thanks in advance. Also, thanks for all the help you as a group have given me when I posted previous questions - sometimes I forget to say thank you. } } Paula. } } Paula Allan-Wojtas } Research Scientist - Food Microstructure } Agriculture and Agri-Food Canada } Atlantic Food and Horticulture Research Centre } Kentville, Nova Scotia Canada B4N 1J5 } } Tel: (902) 679-5566 } FAX: (902) 679-2311 } } email: allanwojtasp-at-em.agr.ca
John Mansfield's reminded me that I would like to have some sort of users' group meeting for Emispec users for mutual support and information exchange. I have discussed this with the Emispec folks, but I don't think that they have followed up on it. Could I seed some level of support for a get-together at M&M MM so that I can send it to Emispec. Please respond to me offline and I will send them the number and names of the people that would desire something like that. I would like to see something in a positive and instructive type of meeting.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center Guys Run Rd. (packages) P. O. Box 11472 (letters) Pittsburgh, PA 15238-0472
this is a message for EM people in Sydney or Woolongong, Australia.
The rotory pump on my JEOL 2020F TEM is short of oil and making a bit of noise so I'm trying to find some quickly. Our JEOL service guys have ordered some "MR100" but it may take a couple of weeks to arrive. Can any one lend me a few hundred mls in the meantime?
Can anyone suggest a local supplier of this rotory pump oil?
Cheers,
Mark Blackford TEM Group Materials Division, ANSTO PMB 1, Menai, N.S.W. Australia 2234
Phone 61 2 9717 3027 Fax 61 2 9543 7179
Disclaimer: The views expressed in this E-mail message do not necessarily represent the official views of ANSTO from which this message was conveyed.
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
My guess would be the current loading at start up. When you turn on an SEM from cold, everything starts and draws current. In particular, the rotary pump kicks in and draws a high current as it starts. You will need to be able to set up the UPS so that it can handle this - or set it on a short timer so that it automatically switches in, say, 10 mins after start up.
Regards, -- Larry Stoter JEOL (UK) Ltd Silver Court, Watchmead, Welwyn Garden City, AL7 1LT, United Kingdom tel: +44-(0)1707-377117, fax: +44-(0)1707-373254, e-mail: larrys-at-jeoleuro.com
Probably you are having problems with the high switch on current the sem draws. You could try switching on different parts of the SEM system not at the same time (if possible). You could also consider to only power the electronics of the SEM via the UPS, in that way you reduce the required current drastically, while there is little use for a pump to sit on the UPS power. Alternatively you could look for a more powerfull UPS that can handle the larger currents (for short moment is enough- see data sheets of different UPS's).
Hope this helps...
Jo
************************************************************* * Jo Verbeeck * * University of Antwerp * * Dept. EMAT (Electron Microscopy for Materials Research) * * e-mail: joverbee-at-ruca.ua.ac.be * * tel: +32(0)3 218 02 49 * * fax: +32(0)3 218 02 57 * *************************************************************
On Tue, 20 Jun 2000, Paula Allan-Wojtas 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, all, } } I am posting this question to see if we can get some help for our building engineer to deal with problems we are having with our SEM (JEOL T330A). } } We suspect that we are having a problem with the building power supply to our SEM. Other folks in our building using other types of equipment have found it necessary to use power conditioners or UPS systems to run their equipment - these systems seem to do the trick for them. } } We are trying this approach with the power supply to our SEM, but find that there are some problems. When using the UPS system, we have to connect it up, put it on bypass, start up the SEM, then switch the UPS over after that - a real nuisance. If we don't do this, the SEM won't start. } } Does anyone else have this problem, and if so, has found an easy solution? Please contact me offline if you have any suggestions and other words of wisdom. Thanks in advance. Also, thanks for all the help you as a group have given me when I posted previous questions - sometimes I forget to say thank you. } } Paula. } } Paula Allan-Wojtas } Research Scientist - Food Microstructure } Agriculture and Agri-Food Canada } Atlantic Food and Horticulture Research Centre } Kentville, Nova Scotia Canada B4N 1J5 } } Tel: (902) 679-5566 } FAX: (902) 679-2311 } } email: allanwojtasp-at-em.agr.ca } } }
If you are using water vapour in your environmental SEM you may want to check the suitability of a Be window. I have a vague memory that water will make holes in the thin (6-8um) Be, check it out with your supplier.
I have used two polymer windows on an EDX detector in a TEM gas reaction cell. The front window is on a replaceable mount in front of the detector. This is easily exchangeable in case I cover it with reaction products from the in-situ experiment; I don't want to see them in all subsequent spectra. Much cheaper then a window replacement and can be carried out by me on site. Contact me for further details if you want to.
Reducing the disturbance to the vacuum system gasses during pumpdown and venting by limiting the speed may help prevent dust particles damaging the window in the first place.
Regards, Ron
On Tue, 20 Jun 2000 tellis2-at-hallmark.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 } -----------------------------------------------------------------------. } } } HI: } For the second year in a row I am having to have our thin window (low } element type-brand name with held) replaced. } We have the EDS connected to a TOPCON SM-500 environmental SEM. I look at a } lot industrial dusts for particle size and composition and the manuf. of } the EDS system says they poke holes in the polymer window. } Has anyone else have this problem? } I have elected to have a thin beryllium installed this time since my } boss is upset about spending $7000 every year plus the downtime and since } low element detection is not critical . } Thanks } Terry Ellis } Hallmark Cards Inc. } } }
=========================================================================== Mr. Ron Doole e-mail ron.doole-at-materials.ox.ac.uk Department of Materials, phone +44 (0) 1865 273701 University of Oxford, fax +44 (0) 1865 283333 Parks Road. Oxford. OX1 3PH. UK. ============================================================================
I am currently studying for an MSc in Textile studies, at Bolton Institute, England.
For part of my studies I have been analysing some polyester film that has been treated with 10% sodium hydroxide under imposed load under polarized light on an optical microscope. Vivid colours have been noted, Reds, greens, and I am struggling to find information to outline what these colours are actually indicating. I therefore write and ask for any information you may deem relevant, I would be extremely grateful for.
Hi, I am working on powders of mixed WC and Co and I have some problems with EDS. EDS analysis on grains whose diffractions patterns are unambiguously indexed in the Co structure give a majority of W, and EDS analysis on grains whose diffractions patterns are unambiguously indexed in the WC structure (without distortion and superstructures) always indicate the presence of Co (with a majority of W). So I was wondering if there could be any problem with EDS on Cobalt (because of magnetism????)
I have an Electroscan 2010 and routinely have problems with the window. I have ascribed it to particles flying around from the surface of the sample. This generally occurs at startup when there may be some charging. We solved the problem by retrofiting the detector so that it can be retracted from the chamber when changing specimens. Its a pain in the butt, but less so than having to replace windows.
Thanks Robert Carlton Aventis Pharmaceuticals robert.carlton-at-aventis.com
Greetings, Yes, those colors are beautiful, arn't they? The colors indicate the magnitude of birefringent retardation in your sample. In essence, you are seeing subtraction colors resulting from the diminution of intensity at certain wavelengths. Say your sample has a retardation of 550 nm. When linearly polarized light of that wavelength passes through the sample, it will be retarded by exactly one wavelength, which is the same as zero retardation. Thus, it will not be affected by the sample and will be blocked by the analyzer. But light of longer or shorter wavelengths will be retarded by more or less than a wave and so will emerge as elliptically polarized light and thus will have a component transmitted through the analyzer. So for any actual sample retardation, the color will result from exactly how much of each wavelength gets through. Because our eyes are very sensitive to color, this has been used for more than 100 years to measure/estimate retardation. There is a chart that reproduces the colors as a function of retardation. I have no idea if this chart has made it on line but I would be careful. The colors are very tricky to print and great care was used to get them right. You would do far better to look this one up in your library. I am sure the Bolton Textile Institute will have excellent books on polarized light microscopy, and while they might be dusty, this particular corner of science has been well understood for years and years. Hope this helps, Tobias Baskin
} } } Dear Sir/Madam } } I am currently studying for an MSc in Textile studies, at Bolton } Institute, England. } } For part of my studies I have been analysing some polyester film that } has been treated with 10% sodium hydroxide under imposed load under } polarized light on an optical microscope. Vivid colours have been noted, } Reds, greens, and I am struggling to find information to outline what } these colours are actually indicating. I therefore write and ask for any } information you may deem relevant, I would be extremely grateful for. } } I thank you for your time. } } Yours Faithfully } } Kelly Goodman } } EMAIL: suite666-at-netscapeonline.co.uk
A colleage of mine here at the Medical College of Wisconsin is working on a new teaching manual for our first-year medical students' cells & tissues course. We are currently putting together a portion of the manual which shows students electron micrographs of cells and organelles. We are looking for high quality TEM images (non-copyrighted) of structures such as the following:
We would be grateful to anyone who feels they have something useful to contribute. The plan is to scan appropriate images for the lab manual and return them as soon as possible to the owner. There will be a list at the end of the manual acknowledging all contributors.
Thank you everyone,
Susan K. Danielson, MS Neuromuscular Lab Coordinator Dept. Neurology, Medical College of Wisconsin ph: 414.259.3836 email: sdaniels-at-mcw.edu
We are looking for qualified professionals to provide applications-based sales and technical support for our complete line of optical microscopes, sample preparation equipment, consumables, and digital imaging systems in the Dallas/Ft. Worth and Austin, TX areas. Must have at least 2 years experience in failure analysis or materials analysis using microscopes and sample preparation equipment. Prior sales experience is not necessary. Understanding of dimensional measurement and image analysis systems a plus.
Please respond by e-mail to mms-at-micrometsys.com
We are looking for qualified professionals to provide applications-based sales and technical support for our complete line of optical microscopes, sample preparation equipment, consumables, and digital imaging systems in the Dallas/Ft. Worth and Austin, TX areas. Must have at least 2 years experience in failure analysis or materials analysis using microscopes and sample preparation equipment. Prior sales experience is not necessary. Understanding of dimensional measurement and image analysis systems a plus.
Please respond by e-mail to mms-at-micrometsys.com
Responding to the message of {3.0.5.32.20000620115226.008c7a80-at-mailserver.aecom.yu.edu} from "David H. Hall" {hall-at-aecom.yu.edu} :
David,
I've done it, but the thing to watch out for is that if the temperature of your acetone/LR Gold mixtures - say 1:1 acetone:LR Gold - drops below about -35 C, some components of the LR Gold will start to freeze out, solutions gets cloudy. For methanol sub-solution's, LR Gold starts to freeze out at even higher temps, about -27 C.
Just experiment with your sub mixtures at various temperatures to see if you get any freeze-out like this happening, before you do an actual sub run. If so just make sure you stay warmer than freezing points.
I'm curious why this happens, anyone else got any info on this?
Gib Ahlstrand
} Does anyone have experience taking specimens from acetone into LR Gold } resin? Some of the brochures on LR White seem to recommend against it, but } we are hoping it may be OK for LR Gold. We are doing freeze substitution } through acetone for EM-immunocytochemistry. } } Thanks in advance for your help. } David H. Hall } Center for C. elegans Anatomy } Department of Neuroscience } 1410 Pelham Parkway } Albert Einstein College of Medicine } Bronx, NY 10461 } } phone (718) 430-2195 FAX (718) 430-8821 } hall-at-aecom.yu.edu } website: www.aecom.yu.edu/wormem
Gib Ahlstrand Electron Optical Facility, University of Minnesota, Dept. Plant Pathology 495 Borlaug Hall, St. Paul, MN. USA. 55108 (612)625-8249 612-625-9728 FAX, giba-at-puccini.cdl.umn.edu http://biosci.umn.edu/MIC/consortium.html
Even though we have not yet had our Microscopy and Microanalysis 2000 meeting in Philadelphia the Program Committee for the M&M 2001 meeting is preparing the symposia topics for the meeting in Long Beach. If anyone has suggestions for topics that they would like to see included in the Long Beach meeting please contact one of the Program Committee Officers listed below. Please note that we may not be able to respond to all suggestions for this year, but voicing your opinion now may get a topic of interest on the program list for upcoming years.
Thanks.
Bob Price
Program Officers:
Bob Price, M&M 2001 Program Chair Price-at-med.sc.edu
Inga Holl Musselman, M&M 2001 MAS Program Co-chair imusselm-at-utdallas.edu
Edgar Voelkl, M&M 2001 Program Vice-chair/M&M 2002 Program Chair vog-at-ornl.gov
Robert L. Price Director, Instrumentation Resource Facility USC School of Medicine Garner's Ferry Road Columbia, SC 29208 Phone: 803-733-3393 Fax:803-733-1533
I couldn't find this in the archives but I cannot imagine someone has not asked this already. If so, please direct me to the approximate month, year. If not, here goes:
We are decomissioning a laboratory refrigerator. Over the 30+ years of use it has accumulated osmium black on the walls. It has been determined by our EH&S people (after consultation with a hazardous waste shipping compnay) that disposal requires a sealed, separate container for the refrigerator, several licenses, and a hazardous waste truck. The estimate is not in yet, but everyone is talking thousands of dollars. If that is the only alternative for safe disposal, so be it. However, I cannot imagine that there is not a safe alternative to decontaminate before shipping. After all we clean up spills with corn oil and reduce osmium tetroxide with 5% sodium bisulfite solution or sodium metabisulfite. Any definitive advice or references will be appreciated.
Thanks-
-- Jay ---------------------------------------------- - AKA: W. Gray Jerome, Ph.D. - - Department of Pathology - - Wake Forest University School of Medicine - - Winston-Salem, NC 27157-1092 - - Ph: 336-716-4972, 336-716-2675 - - Fax: 336-716-6174 - - E-mail: jjerome-at-wfubmc.edu - ----------------------------------------------
W. Gray Jerome wrote: =========================================================== I couldn't find this in the archives but I cannot imagine someone has not asked this already. If so, please direct me to the approximate month, year. If not, here goes:
We are decomissioning a laboratory refrigerator. Over the 30+ years of use it has accumulated osmium black on the walls. It has been determined by our EH&S people (after consultation with a hazardous waste shipping compnay) that disposal requires a sealed, separate container for the refrigerator, several licenses, and a hazardous waste truck. The estimate is not in yet, but everyone is talking thousands of dollars. If that is the only alternative for safe disposal, so be it. However, I cannot imagine that there is not a safe alternative to decontaminate before shipping. After all we clean up spills with corn oil and reduce osmium tetroxide with 5% sodium bisulfite solution or sodium metabisulfite. Any definitive advice or references will be appreciated. ============================================================= I will risk showing what maybe I don't know, but this is really an important question. Laboratories should not be wasting thousands of dollars needlessly, when other alternatives are available.
I thought that Os (IV) oxide, that is, the tetroxide form of osmium, was clear, after all, crystals in ampoules are fairly clear or translucent, and 4% aqueous osmium tetroxide is water clear. And I thought that in the reduced form, that is, the dioxide or Os (II) form, the color was black. Putting it another way, if it is in the reduced form (e.g. black), it is not in its "hazardous" form, and indeed might even be fairly innocuous. A **quick** survey of shipping regulations does not even indicate that osmium (II) oxide is a controlled material from the standpoint of shipping. I want to emphasize the word "quick", but I could not find a UN number asigned for it.
If that is the case, and if I am right, why could not the black deposit be collected, perhaps by way of some rubbing and scrubbing, and disposed of (or better yet, recycled) and the refrigerator disposed of as any ordinary refrigerator might be dealt with?
Remember, I am not saying that this could be done, legally or otherwise, I am just asking what it is that I might be missing here.
Disclaimer: SPI Supplies is a major supplier of osmium tetroxide to EM laboratories worldwide.
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.spi.cc ######################## ============================================
-We need chemical reference standards for the analysis of a Ni-base alloy.
-We intend to use the energy-dispersive X-ray analysis, coupled to a Scanning Electron Microscope(SEM).
-The composition of the alloy is going to vary between 18 to 28 wt% Aluminium ; Al is the most important element to be tested, the remainder of the alloy being mostly Ni of course.
-Secondary main elements are : Cr from 4 to 10 wt% ; Co from 10 to 15 wt% ; Elements in low concentration (below 3%) are Ti and Mo.
-Accuracy : 0.5 % in Al is required.
Questions are :
-How many different standards would we need ?
-What should be the chemical variations from standard to standard to reach the required accuracy ?
-What would be the price for such standards ? Time for delivery ? Where to order this ?
-Can you send us information (reprints from technical journals, articles) on the topic of chemical analysis by means of energy-dispersive (or wavelength-dispersive) X-ray systems ?
If you need any further information, please contact me.
In anticipation, many thanks for a prompt answer.
Yours faithfully,
PAUL-LAURENT CAPRON JR. S.A. LABORIMPEX N.V. RUE DES ALLIES 78-80 BONDGENOTENSTRAAT 78-80 1190 BRUSSEL/BRUXELLES BELGIUM TEL.: 00 32 2 345.99.94 FAX.: 00 32 2 347.39.63
Fellow EM Users, Just a short query ----- would anyone installed a modern day alternative to the 1920's !!! style glass ionization vacuum gauge ("electron current") + electronics box for the JEOL 4B coater? Thankyou for your time, Barry EM UNIT UNSW
Very funny, in every meaning of that word. Of course, the waste company would make a meal of it, why ask?
Its not Os04 that is coating the inside of the fridge, but a tiny amount of osmium metal. The metal is not particularly toxic. Os and Ir are the heaviest metals, with a specific gravity over 22, but nonetheless, its hard to see that you would have 2g of Os in any refrigerator coating. Finely dispersed Os produces tiny amounts of Os04 - if that is a problem, than that would have been so in the confined lab space. If that fridge was dumped conventionally in a land-fill, the tiny amount of OsO4 would be reduced before ever reaching the surface. If it was smelted for scrap, arguably it would, however, immeasurably improve the quality of the scrap. There is an argument for smearing a bit of vegetable oil over the inside of the fridge prior to disposal. Have those safety people given their fully developed reasons? I love to know them.
The bigger picture is interesting too. Safety people are not selected for, nor seem to get taught much logic or a sense of proportion. So we seem to get more and more hassles about things that hardly matter at all, but there is no money and often no objection to the really large environmental issues. Its been said that the old Roman civilasation's collapse was partially due to the difficult Roman numerals, which made the efficient administration of the empire impossible. Our "Western Society" has seen a rapid increase of useless, if not counterproductive rules and regulations, I fear if that trend continues that these "well-meaning" rules will undermine our society. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Thursday, June 22, 2000 7:07 AM, Jay Jerome [SMTP:jjerome-at-wfubmc.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 couldn't find this in the archives but I cannot imagine someone has } not asked this already. If so, please direct me to the approximate } month, year. If not, here goes: } } We are decomissioning a laboratory refrigerator. Over the 30+ years of } use it has accumulated osmium black on the walls. It has been determined } by our EH&S people (after consultation with a hazardous waste shipping } compnay) that disposal requires a sealed, separate container for the } refrigerator, several licenses, and a hazardous waste truck. The } estimate is not in yet, but everyone is talking thousands of dollars. If } that is the only alternative for safe disposal, so be it. However, I } cannot imagine that there is not a safe alternative to decontaminate } before shipping. After all we clean up spills with corn oil and reduce } osmium tetroxide with 5% sodium bisulfite solution or sodium } metabisulfite. Any definitive advice or references will be } appreciated. } } Thanks- } } -- } Jay } ---------------------------------------------- } - AKA: W. Gray Jerome, Ph.D. - } - Department of Pathology - } - Wake Forest University School of Medicine - } - Winston-Salem, NC 27157-1092 - } - Ph: 336-716-4972, 336-716-2675 - } - Fax: 336-716-6174 - } - E-mail: jjerome-at-wfubmc.edu - } ---------------------------------------------- } }
Rotary pump oil is a refined mineral oil, which needs to have low vapour pressure, low viscosity and good lubricating properties. such oils are made by "oil companies". In my lab days I never had the luxury to purchase oil from a microscope manufacturer, because their on-cost for such items are horrific and real and my budgets were small.
Disclaimer: ProSciTech sell Rotary Pump Oil. Don't even ask to ship it to USA; the material is cheap, but expensive to ship. Our price includes shipping within Australia only. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Wednesday, June 21, 2000 2:33 PM, Mark Blackford [SMTP:mgb-at-ansto.gov.au] 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 all, } } this is a message for EM people in Sydney or Woolongong, Australia. } } The rotory pump on my JEOL 2020F TEM is short of oil and making a bit of } noise so I'm trying to find some quickly. Our JEOL service guys have } ordered some "MR100" but it may take a couple of weeks to arrive. Can any } one lend me a few hundred mls in the meantime? } } Can anyone suggest a local supplier of this rotory pump oil? } } Cheers, } } } Mark Blackford } TEM Group } Materials Division, ANSTO } PMB 1, } Menai, N.S.W. } Australia } 2234 } } Phone 61 2 9717 3027 } Fax 61 2 9543 7179 } } Disclaimer: } The views expressed in this E-mail message do not necessarily represent the } official views of ANSTO from which this message was conveyed. } }
I remember using the old Geissler tube on an evaporator 32 years ago and very likely they existed long before then. They were actually a bit of an ongoing maintenance expense, but I loved the pretty colours. Cannot see any reasons why other type gauges could not be used, except the manufacturer would find the Geissler tube a cheaper alternative.
Disclaimer: ProSciTech supplies vacuum gauges, circa year 2000
Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Thursday, June 22, 2000 10:24 AM, Barry Searle [SMTP:B.Searle-at-unsw.edu.au] wrote: } } Fellow EM Users, Just a short query ----- would anyone installed a } modern day alternative to the 1920's !!! style glass ionization vacuum } gauge ("electron current") + electronics box for the JEOL 4B coater? } Thankyou for your time, Barry EM UNIT UNSW }
Regarding the colours in the PET film, the optical part is as Tobias Baskin says. In regard to the polymer science side of it, sretched films or fibres show optical anisotropy, because the polarizability along the polymer chain is different from that transverse to it. The more highly stretched the fibre, the more the chains will be oriented, and this will increase the optical anisotropy of the fibre. Multiply this by the thickness of the fibre and that will give you the birefringence. Most likely you will be seeing coloured fringes parallel to the fibre direction, which indicate the different viewing thickness through the different parts of the fibre, just like the colours in an oil slick under a car indicate different thicknesses of an oil film.
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
I am not familiar with microscopy. The most I know is how SEM/EDX can examine a cross sectional sample.
Currently, I am looking for some methodology to analyse interface structure, elementry and chemical compound especially between the solder and metal, i.e. the IMC layer. The solder height will be ranging from 2-4 mil (SMT process). Wondering anybody can advice the best method to analyse the IMC layer. EDX is capable of detecting the element contain insde the IMC but I am seeing more and more alloy up to ternary or quartenary alloy at the layer. Problem is I do not have any idea about where and how it come from.
I'm looking for a manual for the JVG-N2 ionization vacuum gauge. I've already tried JEOL and they could find JVG-N1 manual (thanks!), but not -N2, and -N1 and -N2 are different enough (different connectors, apparently EB grid heating in -N2 etc.) that the -N1 manual is not enough to get the gauge operational.
So, anyone with manual for JVG-N2 ??
btw: The JVG-N1 manual mentions "Forgel tube" which looks in drawing just like normal hot-filament ionization gauge tube.. Is "Forgel tube" just an old term for one?
For some idea of what can be done, check out the soldering iron tip images and x-ray data at my web site.
Keep in mind that I did not not take time to achieve the best possible polish and most of the work was at rather low magnification.
Site: http://woody.white.home.att.net
Woody White
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Hello,
I am not familiar with microscopy. The most I know is how SEM/EDX can examine a cross sectional sample.
Currently, I am looking for some methodology to analyse interface structure, elementry and chemical compound especially between the solder and metal, i.e. the IMC layer. The solder height will be ranging from 2-4 mil (SMT process). Wondering anybody can advice the best method to analyse the IMC layer. EDX is capable of detecting the element contain insde the IMC but I am seeing more and more alloy up to ternary or quartenary alloy at the layer. Problem is I do not have any idea about where and how it come from.
} question. Laboratories should not be wasting thousands of dollars } needlessly, when other alternatives are available. } } I thought that Os (IV) oxide, that is, the tetroxide form of osmium, was } clear, after all, crystals in ampoules are fairly clear or translucent, and } 4% aqueous osmium tetroxide is water clear. And I thought that in the } reduced form, that is, the dioxide or Os (II) form, the color was black. } Putting it another way, if it is in the reduced form (e.g. black), it is not } in its "hazardous" form, and indeed might even be fairly innocuous. A } **quick** survey of shipping regulations does not even indicate that osmium } (II) oxide is a controlled material from the standpoint of shipping. I want } to emphasize the word "quick", but I could not find a UN number asigned for } it. } } If that is the case, and if I am right, why could not the black deposit be } collected, perhaps by way of some rubbing and scrubbing, and disposed of } (or better yet, recycled) and the refrigerator disposed of as any ordinary } refrigerator might be dealt with? }
The problem is that any form of osmium is a heavy metal, and therefore should be treated as one would cadmium or arsenic. It is not innocuous.
Don }
______________________________________________________________________ Donald L. Lovett e-mail: lovett-at-tcnj.edu Assoc. Professor, Dept. of Biology voice: (609) 771-2876 P.O. Box 7718 fax: (609) 637-5118 The College of New Jersey Ewing, NJ 08628-0718
We have made magnetic nanodots and metastable magnetic phases. The physical structures are well revealed by TEM. However, we would like to relate the magnetic properties with magnetic structure and physical structure. Are there other methods such as neutron diffraction, X-ray or spin polarized electrons that can characterize the magnetic structure/ordering. Please contact me if you could offer such info/collaboration. Thanks a lot.
Yi
******************************************************************* Yi Liu Department of Mechanical Eng. and CMRA 104 N Walter Scott Engineering Center University of Nebraska-Lincoln Lincoln, NE 68588-0656 Tel. (402) 472-7759 (Office) Tel. (402) 472-8762 (EM lab) Fax (402) 472-1465 Email: yliu-at-unlserve.unl.edu *******************************************************************
This is a very complex problem. You don't mention the thickness of the intermetallic layers. EDX analysis has relatively poor spatial resolution in the SEM (of the order of microns) so you can clearly see features that you cannot analyze well. People often use backscatter electron imaging (which can be calibrated) in cases such as yours. You should read a good book on SEM - the subject is too complex to cover in an e-mail like this.
If you can make thin foils of the sample, AEM (analytical electron microscopy) analysis would overcome most of the problems. A LaB6 instrument will give you better than 10 nm analytical resolution, while a field-emitter will approach 1 nm resolution. The problem, though, will be specimen preparation. I would be tempted to suggest using a microtome to cut thin slices, but I don't know the form of your material.
Good luck!
Tony Garratt-Reed
At 06:37 PM 06/22/2000 +0800, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
** Anthony J. Garratt-Reed ** MIT Room # 13-1027 ** 77 Massachusetts Avenue ** Cambridge, MA 02139-4307 ** USA ** ** Phone: (+) 1-617-253-4622 ** Fax: (+) 1-617-258-6479 **
Dear Paul-Laurent, The usual standards that are used for alloy analysis in the SEM+EDX are pure element standards. You can prepare pure (three or four nines, e.g. 99.9 or 99.99%) metal standards yourself, or any of the EM catalog houses will list standards consisting of pure metals or minerals covering the range of most common elements. They are fairly expensive (~$2000US) and I am not sure of delivery time. Tousimis is one standard supplier that I know of. The same standards are used for EDX or WDX analysis. You collect the spectra from these standards under carefully reproducible conditions in your SEM and follow the instructions in your EDX software to do fully standard analysis with matrix corrections. You can store the standard spectra in the EDX computer, so you only need to run them once for each set of conditions. It is quite difficult to get an accurate and precise analysis of Al in Ni alloys, because there is a large correction factor for absorption of the Al x-rays by the Ni matrix, and because the best SEM conditions for Ni analysis, 20 kV, is a large overvoltage for Al. It is very helpful if you can make or obtain a well-characterized, homogeneous alloy in the range of the alloy you will be testing, so you can check the results of your EDX analysis. The last time I did a careful phase analysis of an alloy consisting of 5% Al in Zn, I eventually went to analysing the Al at 10 kV and doing the Zn by difference. This was the only way to get the Al results with the accuracy required and agreeing with the phase diagram. With the other elements present in your system, you cannot use the element-by-difference method, but you could check to see if you get the same Al results at 10 kV, doing the Ni by difference (forgetting the other elements), as you get in your full, standard analysis at 20 kV. Most of the EDX supplier companies have excellent instruction books that cover the topics of chemical analysis by EDX and the exact conditions required for accurate analysis. Some offer courses in EDX analysis and they are a very good way to get familiar with the strengths and weaknesses of the system. Please contact me if you need any more information. At 07:07 PM 6/21/00 -0500, you wrote:
} CAN SOMEBODY HELP ME?: } } -We need chemical reference standards for the analysis of a Ni-base alloy. } } -We intend to use the energy-dispersive X-ray analysis, coupled to a } Scanning } Electron Microscope(SEM). } } -The composition of the alloy is going to vary between 18 to 28 wt% } Aluminium ; } Al is the most important element to be tested, the remainder of the alloy } being } mostly Ni of course. } } -Secondary main elements are : Cr from 4 to 10 wt% ; Co from 10 to 15 } wt% ; Elements in low concentration (below 3%) are Ti and Mo. } } -Accuracy : 0.5 % in Al is required. } } } Questions are : } } -How many different standards would we need ? } } -What should be the chemical variations from standard to standard to reach } the } required accuracy ? } } -What would be the price for such standards ? Time for delivery ? Where to } order this ? } } -Can you send us information (reprints from technical journals, articles) on } the } topic of chemical analysis by means of energy-dispersive (or } wavelength-dispersive) X-ray systems ? } } If you need any further information, please contact me. } } In anticipation, many thanks for a prompt answer. } } Yours faithfully, } } PAUL-LAURENT CAPRON JR.
Regards, Mary
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-interchg.ubc.ca
Hi Listservers, We are in the process of replacing our old critical point dryer and have narrowed our selection to the following choices: Tousimis Samdri 795 semi automatic and the EMS 850 critical point dryers. If anyone out there is familiar with either of these instruments and would care to comment on their advantages and disadvantages, I would appreciate hearing from you. Offline replies are welcome. I will report back with summary of results.
Tom Januszewski Senior Electron Microscopist Molecular and Cellular Imaging Facility UT Southwestern Medical Center at Dallas 5323 Harry Hines Blvd. Dallas, TX 75390-9039 Phone: 214-648-7291 Fax: 214-648-6408 Email: tom.januszewski-at-email.swmed.edu
Dear Kristian, I also have an old JEOL evaporator (JEE-4B) and, yes, the Forgel tube was just a hot-wire ionization gauge. I got tired of buying new ones, so I successfullly replaced that with a cold cathode gauge attached to a tube of the right diameter. At 01:54 PM 6/22/00 +0300, you wrote: } Hello, } } I'm looking for a manual for the JVG-N2 ionization } vacuum gauge. I've already tried JEOL and they could } find JVG-N1 manual (thanks!), but not -N2, and -N1 and } -N2 are different enough (different connectors, apparently } EB grid heating in -N2 etc.) that the -N1 manual is not } enough to get the gauge operational. } } So, anyone with manual for JVG-N2 ?? } } btw: The JVG-N1 manual mentions "Forgel tube" which looks } in drawing just like normal hot-filament ionization } gauge tube.. Is "Forgel tube" just an old term for one? } } Thanks, } } Kristian Ukkonen. Regards, Mary
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-interchg.ubc.ca
This doesn't seem to cover it. Gold and platinum are heavy metals. So are we to treat them as we treat osmium tetroxide? Where does light end and heavy begin anyway? There is a real need here for a proper definition of the risk.
} } The problem is that any form of osmium is a heavy metal, and therefore } should be treated as one would cadmium or arsenic. It is not innocuous. } } Don } } } } ______________________________________________________________________ } Donald L. Lovett e-mail: lovett-at-tcnj.edu } Assoc. Professor, Dept. of Biology voice: (609) 771-2876 } P.O. Box 7718 fax: (609) 637-5118 } The College of New Jersey } Ewing, NJ 08628-0718 } } } }
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Dr Chris Jeffree University of Edinburgh Biological Sciences EM Facility Daniel Rutherford Building King's Buildings EDINBURGH EH9 3JH Tel: +44 (0) 131 650 5345 FAX: +44 (0) 131 650 6563
I received a request for EM autoradiography of C14 labeled polymers. The idea is to trace the distribution of the polymer once it is internalized in the cell. It has been a long while since I have done autoradiography and I was wondering if there are new books and information that is available out there. Is anyone still using this technique. What are the alternatives? I am not too keen on handling radioactive materials again. I would appreciate comments.
******************************************************* Corazon D. Bucana, Ph.D. Department of Cancer Biology U.T. M.D. Anderson Cancer Center 1515 Holcombe Blvd. Box 173 Houston, Texas 77030 Phone: (713) 792-8106 FAX: (713) 792-8747 Email:bucana-at-audumla.mdacc.tmc.edu FAX: (713) 792-8747
Greetings: I am a new subscriber to this list and am looking for input/advice regarding the use of SEM's in undergraduate research. I teach at a liberal arts university with only undergraduate programs in the natural sciences. A geology colleague and myself (I'm a biologist) are exploring possible funding sources which would allow us to purchase an SEM that would be used by biology and geology undergrads in their independent and senior thesis projects. (We currently have a non-functioning ETEC that we would like to replace -- we've been advised that further repair and head-banging would not be cost effective). If anyone is currently engaged in SEM-based research with undergrads, I would like to get your feedback on your experiences (both the pluses and minuses). Although we don't have a TEM, input from those using TEM with undergrads would also be helpful.
Specific Questions:
1) What types of projects are your students pursuing?
2) Are you conducting interdisciplinary projects? e.g., micropaleontology or environmental applications.
3) Are there particular models of SEMs which would be more user friendly since we would be training undergrads and we would also be the primary trainers and technicians?
4) Have you developed an undergrad EM course?
Thanks, Ken
----------------------------------------------------------------------- Kenneth Long, PhD email: long-at-clunet.edu Associate Professor, Biology Office: (805) 493-3346 California Lutheran University Fax: (805) 493-3392 60 West Olsen Rd. #3700 http://www.clunet.edu/Biology/Anatomy Thousand Oaks, CA 91360
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Fellow Microscopists: Stuart Mckernan,M&M'00 Program Chair, has arranged for an additional "EXPERTS" session for the topic of Facility Management. This special session will be held on Wednesday, August 16 at 9:00Am in Room 106. Advertising will be done by flyers, daily news bulletin, and word of mouth.
The room will be available all morning so that we can be somewhat flexible in timing. However, in order to try to make the session as constructive as possible, I would like to identify 2-3 main topics of interest to lab managers from both academic and industrial facilities. These topics will be covered first. Additional topics can then be discussed based on interest and time.
I would appreciate it if those who are interested in attending this session would provide the following: a) 2-3 topics of primary interest to you ranked in order of that interest. b) suggestions of individuals who might be asked to introduce a specific topic so as to give a framework for further discussion.
I am heading out for vacation on July 1 and will unsubscribe from the list a few days prior to then. Please send the above information directly to me rather than to the list so I am sure to get it. We will announce the topics to be discussed and approximate times they will be discussed so that you can choose when you want to attend the session.
If the session is well attended, then it may be added to the agendas of future meeting. Hope to hear from many of you regarding the topics. Debby
Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
by ultra5.microscopy.com (8.9.1b+Sun/8.9.1) id SAA15867 for dist-Microscopy; Thu, 22 Jun 2000 18:00:09 -0500 (CDT) Received: from no_more_spam.com (sparc5 [206.69.208.10]) by ultra5.microscopy.com (8.9.1b+Sun/8.9.1) with SMTP id RAA15863 for "MicroscopyFilteredEmail-at-msa.microscopy.com"; Thu, 22 Jun 2000 17:59:39 -0500 (CDT) Received: from pdx.edu (tuttle.oit.pdx.edu [131.252.120.29]) by ultra5.microscopy.com (8.9.1b+Sun/8.9.1) with ESMTP id RAA15856 for {microscopy-at-sparc5.microscopy.com} ; Thu, 22 Jun 2000 17:59:28 -0500 (CDT) Received: from [131.252.192.235] (host-192-235.dhcp.pdx.edu [131.252.192.235]) by pdx.edu (8.9.3/8.9.3) with ESMTP id PAA26977 for {microscopy-at-sparc5.microscopy.com} ; Thu, 22 Jun 2000 15:53:53 -0700 (PDT) Message-Id: {200006222253.PAA26977-at-pdx.edu} X-Mailer: Microsoft Outlook Express Macintosh Edition - 4.5 (0410)
Hi everyone,
I'm currently making 1.5 litre batches of 4% formaldehyde using paraformaldehyde, sodium hydroxide and distilled water and not having much success. The mixture immediately begins to polymerize. The recipe I used is simply an amplified version of Karnovsky's recipe for 25ml of 4% (which works perfectly). I am wondering whether my assumption of a simple linear relationship between the proportions of NaOH and paraformaldehyde is incorrect. Making stock solutions of higher concentrations does not work either.
Does anyone know a more successful method of preparing large volumes of formaldehyde? We do not want to use phosphates or preservatives.
regards Elizabeth McKenzie -------------------------------------------------- Geomicrobiology and Electron Microscopy Laboratory Room S9 Cramer Hall 1721 SW Broadway Portland State University Portland OR97201
Fellow EM Users, Would anyone have a list of the O rings for use on the JEOL JEE4B/4C vacuum evaporator. The photocopy of pages from the manual that I have does not list any o-rings. Thankyou Barry EM UNIT UNSW
One response. No follow through. Board is going into the circular file.
gg
} Date: Thu, 15 Jun 2000 07:55:38 -0700 } To: MSA listserver } From: "Dr. Gary Gaugler" {gary-at-gaugler.com} } Subject: Polaron E5200 control board } } I have a control PC board for the E5200 sputter coater. } This is the model with an Intel single chip MPU on one } end and a 4-conductor socket on the other end. The } board uses a VME connector for main interface. } } Coater is trashed. Board is OK. If anybody can use } the board, first request gets it. } } gary g.
I had a look at the excellent site http://www.webelements.com/index.html
clicking on osmium and then asking for "biology" this info is given: "Osmium metal does not normally cause problems as it is relatively unreactive but all osmium compounds should be regarded as highly toxic. The metal dust is an irritant and presents a fire and explosion hazard. Osmium oxide, OsO4, is highly toxic, and boils at 130?C (760 mm). Concentrations in air as low as 10-7 g m-3 can cause lung congestion, skin damage, and severe eye damage. The oxide, in particular, should only ever be handled by a properly qualified chemist." We all know about the tetroxide, but the point I made in my previous email was that the fridge contamination is Osmium metal and this has little reactivity. Osmium when finely dispersed would produce a tiny amount of tetroxide. Possibly not enough to worry while in the lab, certainly not if its in landfill or smelted. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Ph +61 7 4774 0370 Fax:+61 7 4789 2313 service-at-proscitech.com Great microscopy catalogue, 500 Links, MSDS, User Notes www.proscitech.com
On Friday, June 23, 2000 6:58 AM, Chris Jeffree [SMTP:cjeffree-at-srv0.bio.ed.ac.uk] wrote:} } } This doesn't seem to cover it. Gold and platinum are heavy metals. } So are we to treat them as we treat osmium tetroxide? } Where does light end and heavy begin anyway? There is a real } need here for a proper definition of the risk. } } } } } The problem is that any form of osmium is a heavy metal, and therefore } } should be treated as one would cadmium or arsenic. It is not innocuous. } } } } Don } } } } } } } ______________________________________________________________________ } } Donald L. Lovett e-mail: lovett-at-tcnj.edu } } Assoc. Professor, Dept. of Biology voice: (609) 771-2876 } } P.O. Box 7718 fax: (609) 637-5118 } } The College of New Jersey } } Ewing, NJ 08628-0718 } } } } } } } } } } } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ } Dr Chris Jeffree } University of Edinburgh } Biological Sciences EM Facility } Daniel Rutherford Building } King's Buildings EDINBURGH EH9 3JH } Tel: +44 (0) 131 650 5345 } FAX: +44 (0) 131 650 6563 } } Inveresk Cottage, 26 Carberry Road, } Inveresk, Musselburgh, Midlothian EH21 8PR, UK } Tel. +44 (0) 131 665 6062 / Mobile 0410 585 401 } ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Dear colleagues, Many thanks for all responses to my question considering the hole in the EDX spectrum. We had a service engineer in our lab and he tried to solve the problem but he was not successful. I have prepared WEB page with the images of spectra, that were recorded from following samples: Cu_Al sample (usually used for calibration) Ti grid AL grid K standard
The url of that page is following: http://www.biomed.cas.cz/~benada/EDX_page/index.htm
Please, if anyone would be so kind and could comment the spectra, we will be very happy. Best regards from Prague Oldrich Benada
+-----------------------------------+ Oldrich Benada Acad. Sci. CR Institute of Microbiology Laboratory of electron microscopy Videnska 1083 CZ - 142 20 Prague 4 - Krc Czech Republic +------------------------------------+ Phone: +420-2-4752399 Fax: +420-2-4715743 WEB:http://www.biomed.cas.cz/mbu/lem113/lem.htm
There's nothing mysterious about this. It should be possible to make any volume of formaldehyde solution over a wide range of concentrations. We routinely make 10% solutions without problems in volumes much greater than 25ml. There is no reason to suppose that the volume is relevant, and the concentration is not critical. The temperature needs to be } 60oC and the solution needs to be slightly alkaline. Otherwise it is extremely straightforward. Could you tell us your procedure, step by step? Then maybe it will be possible to diagnose the problem. Chris
Date sent: Thu, 22 Jun 2000 15:50:11 -0700
Commerially available formaldehyde contains formic acid and methanol and is therefore unsuitable for EM. Instead, one usually prepares a solution of formaldehyde from paraformaldehyde. An aqueous solution (40%, w/v) can be made by warming for 1 h at 65ˇV (constant stirring). Milkyness can be overcome by adding a few drops of a 40% (w/v) NaOH solution. This solution is stable for several weeks at 4ˇc (in the dark). Hopes this helps.
De Pauw Bart Ghent University Faculty of Veterinary Medicine Department Morphology
"E. J. McKenzie" schreef:
} ------------------------------------------------------------------------ } 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 everyone, } } I'm currently making 1.5 litre batches of 4% formaldehyde } using paraformaldehyde, sodium hydroxide and distilled water and not } having much success. The mixture immediately begins to polymerize. The } recipe I used is simply an amplified version of Karnovsky's recipe for 25ml } of 4% (which works perfectly). I am wondering whether my assumption of a } simple linear relationship between the proportions of NaOH and } paraformaldehyde is incorrect. Making stock solutions of higher } concentrations } does not work either. } } Does anyone know a more successful method of preparing large } volumes of formaldehyde? We do not want to use phosphates or } preservatives. } } regards } Elizabeth McKenzie } -------------------------------------------------- } Geomicrobiology and Electron Microscopy Laboratory } Room S9 Cramer Hall } 1721 SW Broadway } Portland State University } Portland } OR97201 } } ph:503 725 3362 } fax:503 725 3025
Here's an interesting problem. The other day I was examining some plankton trap samples in our ESEM, and found a few bits of what I took to be pieces of foraminiferal shell - they were slightly curved, crystalline-looking shards with closely spaced fine openings (the foramen, for which forams are named) and a few large holes (possible spine attachment localities). I ran EDS on them to make sure that is indeed, what they were. Planktic foram shells should be fairly pure calcium carbonate. What I got instead were spectra each showing a fairly large strontium peak, a smaller sulfur peak and a little oxygen. There were also very high aluminum peaks on each one, but since the sample was simply air dried on an aluminum stub, I assume that was the source of the Al. In 20-some years of micropaleontology I've never heard of any marine organism that uses strontium to build its shell. Sr is just below Ca in the periodic table, so I suppose it shares some of the same reactability characteristics (but I'm no chemist), so perhaps it could substitute, if no Ca was available (which would be darn funny in a surface marine sample). Let me add that all the other peaks were in their proper places, and the instrument had been calibrated quite recently, and I can think of no source of strontium contamination which could have gotten in there. We're planning on sending some of these bits out for analysis by some other means just to make sure, but for the time being, it's quite a mystery. Any ideas from the marine biologists? The chemists? The EDS gurus? Psychics?
F.C. Thomas MicroAnalysis Facility Geological Survey of Canada (Atlantic) Bedford Institute of Oceanography Dartmouth, Nova Scotia Canada B2Y 4A2
We use the Agfa DD3700 paper processor. However, the wonderful graded papers are no longer available and have not been for some time now. Our back-up system is the Mohr Pro-8. Most of our older trained users are unwilling to use the AGFA multi-contrast graded papers, stating that the iamge is "not as crisp, not as white, harder to work with", etc. If anyone has used AGFA graded papers in the past, and has found a resin-coated graded paper alternative that works to give sharp, crisp, clean, white, micrographs from the DURST Point source, I would like to hear about your solutions. Some un-tried recommendations have been: FOMA papers, FORTE papers, ILFORD papers, and then there is the rest of the list, EFKE, Kentmere, Kodak, MACO, Oriental, adn ECCO. Does anyone know if there is a graded paper comparable to the AGFA RAPITONE GRADES P1,P2, P 3, and P4. AGFA only suggests to use their polycontrast resin papers. I have swithced to digital, but the older folks with all the great publications refuse to lower their standards. I would lke to know more about how I can get the wanted details back onto my micrographs. ANY IDEAS ? ....-TOM
Thomas A Baginski Technical Coordinator for Microscopy Uniformed Services University of the Health Sciences Bethesda, MD 20814-4799
Depending on the polymorph of CaCO3 that foram shells are made of, a few hundred (calcite) to a few thousand (aragonite) ppm of Sr would not be unusual in carbonate precipitated from standard sea-water. SO4 ion is also a fairly significant component of sea-water and S can be incorporated in the structures of carbonate minerals in small amounts. It could also be present as fluid inclusions. Another possibility is that the seawater present in the pores of the forams on recovery from the ocean has precipitated celestite (SrSO4) on dessication, in which case the Sr might have a very patchy distribution.
Roger Mason
Frank Thomas 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 } -----------------------------------------------------------------------. } } Here's an interesting problem. The other day I was examining some plankton } trap samples in our ESEM, and found a few bits of what I took to be pieces } of foraminiferal shell - they were slightly curved, crystalline-looking } shards with closely spaced fine openings (the foramen, for which forams are } named) and a few large holes (possible spine attachment localities). I ran } EDS on them to make sure that is indeed, what they were. Planktic foram } shells should be fairly pure calcium carbonate. What I got instead were } spectra each showing a fairly large strontium peak, a smaller sulfur peak } and a little oxygen. There were also very high aluminum peaks on each one, } but since the sample was simply air dried on an aluminum stub, I assume } that was the source of the Al. }
} We use the Agfa DD3700 paper processor. However, the wonderful } graded papers are no longer available and have not been for some time now. } Our back-up system is the Mohr Pro-8. Most of our older trained users are } unwilling to use the AGFA multi-contrast graded papers, stating that the } iamge is "not as crisp, not as white, harder to work with", etc. } If anyone has used AGFA graded papers in the past, and has found a } resin-coated graded paper alternative that works to give sharp, crisp, clean, } white, micrographs from the DURST Point source, I would like to hear about } your solutions. } Some un-tried recommendations have been: FOMA papers, FORTE papers, } ILFORD papers, and then there is the rest of the list, EFKE, Kentmere, } Kodak, MACO, Oriental, adn ECCO. Does anyone know if there is a graded } paper comparable to the AGFA RAPITONE GRADES P1,P2, P 3, and P4. AGFA } only suggests to use their polycontrast resin papers. } I have swithced to digital, but the older folks with all the great } publications refuse to lower their standards. } I would lke to know more about how I can get the wanted details } back onto my micrographs. ANY IDEAS ? ....-TOM } } Thomas A Baginski } Technical Coordinator for Microscopy } Uniformed Services University of the Health Sciences } Bethesda, MD 20814-4799 } } Voice Phone: 301 295 5691 } Fax: 301 319 8218 } Email: tbaginski-at-usuhs.mil
Dear Tom:
I have had good results with both Kodak and Ilford multigrade resin coated papers in my DD-3700. I am surprised that you can see a difference in sharpness between modern. glossy papers. Is Kodabrom RC still available in grades?
Geoff -- ********************************************** 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 **********************************************
Frank, I am not a microbiologist or even close - I work in cathodoluminescence instrumentation. I recalled that Sheldon Sommer made some comments on Sr in shells in his work at Penn State. On page 282 of his paper, he talks about 70 mole percent strontianite in pelecypods, for what it is worth. Sommer, S. E. CL of carbonates,2. Geological Applications. Chemical geology, 1972, pp. 275-284.
Don Marshall
Donald J. Marshall Relion Industries P.O. Box 12 Bedford, MA 01730 Ph: 781-275-4695 FAX: 781-271-0252 email dmrelion-at-world.std.com
Cathodoluminescence, mass spectroscopy, electron beam technology
"A weed is a flower out of place."
(Please note: Do not send email with attachments to this address. Instead, send it to donbarlen-at-aol.com. Thank you.)
There is a group of marine protistans that uses strontium sulphate to make their tests. Amoeboid critters closely related to ... wait a bit while I dredge through my memory here, my texts are at home ... Actinaria? Relatives of the Radiolarians and Heliozoa IF I'm remembering correctly. They're mentioned inter alia in the book "Synoptic Classification of Living Organisms" and any good invertebrate zoology text or Protistology text.
Phil The Al is most likely from the stub ... try a carbon mount.
} Here's an interesting problem. The other day I was examining } some plankton } trap samples in our ESEM, and found a few bits of what I took to be pieces } of foraminiferal shell - they were slightly curved, crystalline-looking } shards with closely spaced fine openings (the foramen, for which forams are } named) and a few large holes (possible spine attachment localities). I ran } EDS on them to make sure that is indeed, what they were. Planktic foram } shells should be fairly pure calcium carbonate. What I got instead were } spectra each showing a fairly large strontium peak, a smaller sulfur peak } and a little oxygen. There were also very high aluminum peaks on each one, } but since the sample was simply air dried on an aluminum stub, I assume } that was the source of the Al. } In 20-some years of micropaleontology I've never heard of any marine } organism that uses strontium to build its shell. Sr is just below Ca in the } periodic table, so I suppose it shares some of the same reactability } characteristics (but I'm no chemist), so perhaps it could substitute, if no } Ca was available (which would be darn funny in a surface marine sample). } Let me add that all the other peaks were in their proper } places, and the } instrument had been calibrated quite recently, and I can think of no source } of strontium contamination which could have gotten in there. We're planning } on sending some of these bits out for analysis by some other means just to } make sure, but for the time being, it's quite a mystery. } Any ideas from the marine biologists? The chemists? The EDS gurus? } Psychics? } } } F.C. Thomas } MicroAnalysis Facility } Geological Survey of Canada (Atlantic) } Bedford Institute of Oceanography } Dartmouth, Nova Scotia } Canada B2Y 4A2
-- }}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{ Philip Oshel Supervisor, AMFSC and BBPIC Dept. of Animal Health and Biomedical Sciences University of Wisconsin 1656 Linden Drive Madison, WI 53706-1581 voice: (608) 263-4162 fax: (608) 262-7420 (dept. fax)
Below is the result of your feedback form. It was submitted by (barbarac-at-biols.susx.ac.uk) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Friday, June 23, 2000 at 08:00:37 ---------------------------------------------------------------------------
Email: barbarac-at-biols.susx.ac.uk Name: Barbara Ciani School: University of Sussex Question: Hi everyone,
I need to do some congo red staining on my peptides and i need to make sure they stick on the microscope slides.
I have been suggested to use gelatine with chrome alum but my slides don't seem to get 'sticky'.
Any suggestion?
I use 0.5% gelatine (300 bloom) and 0.05% chrome alum.
Date sent: Mon, 19 Jun 2000 09:46:49 -0400 To: {Microscopy-at-sparc5.microscopy.com} } From: Leona Cohen-Gould {lcgould-at-mail.med.cornell.edu}
Tom,
Although we do very little photographic printing anymore, I printed extensively in the past and have gotten excellent results from Kodak and Ilford multi-contrast resin-coated papers, using a Durst professional enlarger, as well as Beseler enlargers of various types. What I have never done, though, is make much use of a paper processor. Force of habit, maybe, but I always preferred tray processing, even when I had to make lots of prints in short amounts of time.
Even resin-coated papers respond somewhat to variations in developing times/temperatures, etc., allowing minor adjustments during processing. Often it is possible to watch the image come up in the tray and know within 15-30 seconds whether it will acceptable or what changes will be necessary. I can only speak for myself, but quality-wise I believe that tray processing is superior, in terms of tonality, grey-scale range, etc. I also believe it's as fast or faster.
That said, I know that people accustomed to processors are just as stubborn as we dinosaurs who prefer tray processing, and nobody is likely to change long-established habits. Any mainstream resin-coated, glossy paper should capture any needed detail that's present in the negative. I'm not saying that photo paper is as "sharp" in terms of lines/mm as a TEM negative, but only that the detail it won't capture is too small to see in normal viewing, and certainly won't survive the reproduction process in a journal. If you're visibly losing detail in the print at a standard, or even close, viewing distance without a lens, I would be checking the negatives and enlarger.
I won't even get into the debate about journals considering digital imaging as a "lowering of standards"! Not going there... :-)
Just my very subjective two cents.
All the best, Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
-----Original Message----- } From: Thomas A. Baginski [mailto:tombg-at-bictom.usuf1.usuhs.mil] Sent: Friday, June 23, 2000 7:09 AM To: Microscopy-at-sparc5.microscopy.com
We use the Agfa DD3700 paper processor. However, the wonderful graded papers are no longer available and have not been for some time now.
Our back-up system is the Mohr Pro-8. Most of our older trained users are
unwilling to use the AGFA multi-contrast graded papers, stating that the iamge is "not as crisp, not as white, harder to work with", etc. If anyone has used AGFA graded papers in the past, and has found a resin-coated graded paper alternative that works to give sharp, crisp, clean, white, micrographs from the DURST Point source, I would like to hear about
your solutions. Some un-tried recommendations have been: FOMA papers, FORTE papers,
ILFORD papers, and then there is the rest of the list, EFKE, Kentmere, Kodak, MACO, Oriental, adn ECCO. Does anyone know if there is a graded paper comparable to the AGFA RAPITONE GRADES P1,P2, P 3, and P4. AGFA only suggests to use their polycontrast resin papers. I have swithced to digital, but the older folks with all the great publications refuse to lower their standards. I would lke to know more about how I can get the wanted details back onto my micrographs. ANY IDEAS ? ....-TOM
Thomas A Baginski Technical Coordinator for Microscopy Uniformed Services University of the Health Sciences Bethesda, MD 20814-4799
} Dear all } } I just found some cells with granules resembling neurosecretory granules in } invertebrate tissues, and I need to confirm their nature. } } Can anyone recomend a straightforward and reliable method to identify } neurosecretory cells (at the light or/and electron microscopic levels)? } } Thanks to all } Dr. A.P. Alves de Matos
Chrome-alum hematoxylin or Falhmi's aldehyde fuchsin work well with mammalian tissues. Check almost any edition of Humason's "Animal Tissue Techniques" for specifics. Modifications for invert tissues are out there but I don't have references handy.
Geoff -- ********************************************** 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 **********************************************
Hi Y'all: Thanks for your overwelming response to my request for an independent FIB lab that can do TEM x-sections. Over the years, this forum has been an invaluable resource for me. Regards, Mike Coviello Lab Manager University of Texas -at- Arlington
Hello everyone, I'm studying microstructures and intergrowths of fibrous minerals by TEM. I'm looking for a sample of amosite and a sample of antophillite fibres in a rock matrix in order to continue this study. I'm gratefully if someone will send my these samples and will be interested to collaborate.
Thank you everyone, Elena Belluso
---------------------------------------------------- Elena BELLUSO Dipartimento di Scienze Mineralogiche e Petrologiche Via Valperga Caluso, 35 I-10125 TORINO - ITALIA tel:(39) 011.670.71.35 - fax: (39) 011.670.71.28 e-mail: belluso-at-dsmp.unito.it http://www.dsmp.unito.it ----------------------------------------------------
The deadline for contributions to the "LATE-BREAKING POSTER SESSION" at MICROSCOPY AND MICROANALYSIS 2000 is very close at hand.
This poster session will be composed of presentations of newly acquired data or analyses which were unavailable for submission by the February 15 deadline. A short, half page abstract describing the studies is required. The abstract should include: Title, Authors, Authors affiliation, and a Brief Description of the studies. The description should include the Aim of the studies, a short characterization of the Methods, and a brief account of the Results and their Importance. The abstracts will obviously not be published in the proceedings, since it is already being printed, but will be available at the meeting. The posters will be advertized in the daily meeting newsletters to alert attendees to their presence.
Abstracts should be e-mailed or faxed to the program chair, Stuart McKernan, at stuartm-at-tc.umn.edu (email) or 612-625-5368 (fax). Abstracts must be received by June 23, 2000. Abstracts will be reviewed by members of the program committee. A limited number of poster boards are available and preference will be given to early submissions. Abstract authors will be notified of acceptance of their abstracts no later than July 1 (earlier for early submissions).
It is not too early to register for the meeting, the pre-meeting congress, or the short courses (or any combination of these events). The early registration deadline is also fast approaching, and can be done using the online registration site at: http://www.peregrine.net/mm2000/
Looking forward to a great meeting in Philadelphia,
__________________ Stuart McKernan stuartm-at-tc.umn.edu Director Office:(612) 626-7594 IT Characterization Facility, University of Minnesota Desk:(612) 624-6009 100 Union Street S. E., Minneapolis, MN 55455 NEW-} Fax:(612) 625-5368
Hello everyone, I'm studying microstructures and intergrowths of fibrous minerals by TEM. I'm looking for a sample of amosite and a sample of antophillite fibres in a rock matrix in order to continue this study. I'm gratefully if someone will send my these samples and will be interested to collaborate.
Thank you everyone, Elena Belluso
---------------------------------------------------- Elena BELLUSO Dipartimento di Scienze Mineralogiche e Petrologiche Via Valperga Caluso, 35 I-10125 TORINO - ITALIA tel:(39) 011.670.71.35 - fax: (39) 011.670.71.28 e-mail: belluso-at-dsmp.unito.it http://www.dsmp.unito.it ----------------------------------------------------
Yes...very interesting result! My guess is that you have happened upon a very rare group of microplankton which make a celestite test (SrSO4). If I remember my micropaleo, look for a group called Acantharia (?) Try Haq & Boersma's "Marine Micropaleontology" for a start. The reason you have not come across them in 20-odd years is that the celestite tests are very unstable...I saw a few beautiful examples in filtered water from the Arabian Sea, never in sediment samples. So not much use for paleoclimatology, but an interesting diversion. If you can post some pics to a website, I'd like to take a look at them.
Matt
Matthew J. Lynn, Ph.D. Center for Advanced Microscopy University of Miami (305)284-4736 mlynn-at-miami.edu
On Friday, June 23, 2000 6:59 AM, Frank Thomas [SMTP:thomasf-at-AGC.BIO.NS.CA] 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 } -----------------------------------------------------------------------. } } } Here's an interesting problem. The other day I was examining some plankton } trap samples in our ESEM, and found a few bits of what I took to be pieces } of foraminiferal shell - they were slightly curved, crystalline-looking } shards with closely spaced fine openings (the foramen, for which forams are } named) and a few large holes (possible spine attachment localities). I ran } EDS on them to make sure that is indeed, what they were. Planktic foram } shells should be fairly pure calcium carbonate. What I got instead were } spectra each showing a fairly large strontium peak, a smaller sulfur peak } and a little oxygen. There were also very high aluminum peaks on each one, } but since the sample was simply air dried on an aluminum stub, I assume } that was the source of the Al. } In 20-some years of micropaleontology I've never heard of any marine } organism that uses strontium to build its shell. Sr is just below Ca in the } periodic table, so I suppose it shares some of the same reactability } characteristics (but I'm no chemist), so perhaps it could substitute, if no } Ca was available (which would be darn funny in a surface marine sample). } Let me add that all the other peaks were in their proper places, and the } instrument had been calibrated quite recently, and I can think of no source } of strontium contamination which could have gotten in there. We're planning } on sending some of these bits out for analysis by some other means just to } make sure, but for the time being, it's quite a mystery. } Any ideas from the marine biologists? The chemists? The EDS gurus? } Psychics? } } } F.C. Thomas } MicroAnalysis Facility } Geological Survey of Canada (Atlantic) } Bedford Institute of Oceanography } Dartmouth, Nova Scotia } Canada B2Y 4A2
I think you have massive icing of your detector crystal, or equally massive contamination of your window. I have modelled your Al spectrum (using DTSA) and can get quite good agreement with your spectrum by putting in 300 microns of ice in the model. The low energy rise is probably caused simply by the tail of the electronic noise which is amplified so much by the scale at which you display your spectrum. Because of the way x-ray absorption works, I can't really tell the difference between absorption in ice and absorption in oil. Anyway, fixing an icing problem is far easier than fixing an oil problem - you simply have to warm up the detector while pumping on it. This will need an adaptor to fit on the pumping port of your detector - your service engineer should have, or be able to get hold of, one of these. So I would try warming the detector and seeing of it solved the problem.
I wouldn't stake my professional reputation on this diagnosis, but I am certain that if the problem is not contamination, then it is a geometrical problem - the collimator may have moved, or some other mechanical derangement taken place.
Good luck,
Tony Garratt-Reed.
At 11:11 AM 06/23/2000 +0200, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
** Anthony J. Garratt-Reed ** MIT Room # 13-1027 ** 77 Massachusetts Avenue ** Cambridge, MA 02139-4307 ** USA ** ** Phone: (+) 1-617-253-4622 ** Fax: (+) 1-617-258-6479 **
Dear Ken, I have always run undergraduate courses on the SEM/EDX and we currently run a course that puts groups of second year engineering students doing independent research projects. They research the composition of many common household and commercial appliances, taking them apart to see what metals, plastics and ceramics are used for the various applications such as small electric motors, heating elements and temperature control. The students are very appreciative of being allowed to sit down and use the instrument with minimal supervision and I find they are very careful with it. I run courses for Anthropology and Electrical Engineering and all disciplines of Engineering, Physics and Chemistry use the instrument for both undergraduate and graduate research. I would suggest you get an EDX and a low vacuum ("environmental") SEM for full versatility in both physical and biological EM. All of my EM instruments (two SEMs with EDX, one 200 kV TEM and SEM with EBSP) are Hitachi, as I have found them the most robust. They run for years of student use and nothing ever seems to go wrong with them. I had an ETEC before my first Hitachi SEM and I liked it too, but they have been out of business for a long time. We also have a fourth year course that covers XRD, SEM, EDX and TEM/EDX. The professor in charge has developed the lab course to touch all of the important principles and demonstrate them in the lab. We cover changing kV, EDX excitation volume, EDX matrix corrections, TEM use and TEM/EDX issues. Please contact me if I can give you any other information. At 03:05 PM 6/22/00 -0700, you wrote: } } Greetings: } I am a new subscriber to this list and am looking for input/advice } regarding the use of SEM's in undergraduate research. I teach at a liberal } arts university with only undergraduate programs in the natural sciences. } A geology colleague and myself (I'm a biologist) are exploring possible } funding sources which would allow us to purchase an SEM that would be used } by biology and geology undergrads in their independent and senior thesis } projects. (We currently have a non-functioning ETEC that we would like to } replace -- we've been advised that further repair and head-banging would } not be cost effective). If anyone is currently engaged in SEM-based } research with undergrads, I would like to get your feedback on your } experiences (both the pluses and minuses). Although we don't have a TEM, } input from those using TEM with undergrads would also be helpful. } } Specific Questions: } } 1) What types of projects are your students pursuing? } } 2) Are you conducting interdisciplinary projects? e.g., micropaleontology } or environmental applications. } } 3) Are there particular models of SEMs which would be more user friendly } since we would be training undergrads and we would also be the primary } trainers and technicians? } } 4) Have you developed an undergrad EM course? } } Thanks, } Ken
Regards, Mary
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-interchg.ubc.ca
Date sent: Fri, 23 Jun 2000 11:57:42 -0400 (EDT) } From: Donald Lovett {lovett-at-TCNJ.EDU} To: Chris Jeffree {cjeffree-at-srv0.bio.ed.ac.uk}
When AGFA (Bayer) pulled the plug on the graded papers, we switched to Kodak Polycontrast RCIII. So I pulled the plug on their activator and fixer and now use Kodak Polymax developer 1:4 dilution and Rapid Fixer in my DD3700. Still use my Durst S45EM with point light source and Kodak Polycontrast Filter Kit. Bob Santoianni Emory University Hospital Atlanta, GA
} Here's an interesting problem. The other day I was examining some } plankton } trap samples in our ESEM, and found a few bits of what I took to be pieces } of foraminiferal shell - they were slightly curved, crystalline-looking } shards with closely spaced fine openings (the foramen, for which forams are } named) and a few large holes (possible spine attachment localities). I ran } EDS on them to make sure that is indeed, what they were. Planktic foram } shells should be fairly pure calcium carbonate. What I got instead were } spectra each showing a fairly large strontium peak, a smaller sulfur peak } and a little oxygen. There were also very high aluminum peaks on each one, } but since the sample was simply air dried on an aluminum stub, I assume } that was the source of the Al. } In 20-some years of micropaleontology I've never heard of any marine } organism that uses strontium to build its shell. Sr is just below Ca in the } periodic table, so I suppose it shares some of the same reactability } characteristics (but I'm no chemist), so perhaps it could substitute, if no } Ca was available (which would be darn funny in a surface marine sample). } Let me add that all the other peaks were in their proper places, } and the } instrument had been calibrated quite recently, and I can think of no source } of strontium contamination which could have gotten in there. We're planning } on sending some of these bits out for analysis by some other means just to } make sure, but for the time being, it's quite a mystery. } Any ideas from the marine biologists? The chemists? The EDS gurus? } Psychics?
Frank -
You've just rediscovered the strontianate Radiolaria. They DO selectively concentrate strontium. I almost selected them as my thesis topic 'way back in the dark ages; I dropped the idea when I realised that there was no way to culture them in the lab. I don't think there is, yet. } Caroline
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
This may be a bit of a shameless plug for a paper I contributed to, but here's the reference:
Payne CM, Cromey DW (1987) Ultrastructural similarities between Chlorohydra viridissima and human neurosecretory granules: A cytochemical study using the uranaffin reaction. Cytobios 50:191-203.
Back when I was doing clinical TEM, we used the Uranaffin Method as the "gold standard" for diagnostic identification of neurosecretory granules in tumors. It was more reliable than immunomarkers and its fairly simple to do. The reference section from this particular article should include Dr. Payne's original publication on the technique. The most important thing is to follow the protocol, failure to use the specified buffers usually causes problems.
Yours, Doug Cromey
At 09:46 AM 6/23/2000 -0400, you wrote: } "A.P. Alves de Matos" wrote: } } I just found some cells with granules resembling neurosecretory granules in } } invertebrate tissues, and I need to confirm their nature. } } } } Can anyone recomend a straightforward and reliable method to identify } } neurosecretory cells (at the light or/and electron microscopic levels)? } } Chrome-alum hematoxylin or Falhmi's aldehyde fuchsin work well with mammalian } tissues. Check almost any edition of Humason's "Animal Tissue Techniques" for } specifics. Modifications for invert tissues are out there but I don't have } references handy.
.................................................................... : Douglas W. Cromey, M.S. Dept. of Cell Biology & Anatomy : : Research Specialist, Principal University of Arizona : : (office: AHSC 4212A) P.O. Box 245044 : : (voice: 520-626-2824) Tucson, AZ 85724-5044 USA : : (FAX: 520-626-2097) (email: doug-cromey-at-ns.arizona.edu): :...................................................................: http://www.pharmacy.arizona.edu/exp_path.html Home of: "Microscopy and Imaging Resources on the WWW"
Sorry if this is double-posted....server troubles!
Frank,
Yes...very interesting result! My guess is that you have happened upon a very rare group of microplankton which make a celestite test (SrSO4). If I remember my micropaleo, look for a group called Acantharia (?) Try Haq & Boersma's "Marine Micropaleontology" for a start. The reason you have not come across them in 20-odd years is that the celestite tests are very unstable...I saw a few beautiful examples in filtered water from the Arabian Sea, never in sediment samples. So not much use for paleoclimatology, but an interesting diversion. If you can post some pics to a website, I'd like to take a look at them.
Matt
On Friday, June 23, 2000 6:59 AM, Frank Thomas [SMTP:thomasf-at-AGC.BIO.NS.CA] 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 } -----------------------------------------------------------------------. } } } Here's an interesting problem. The other day I was examining some plankton } trap samples in our ESEM, and found a few bits of what I took to be pieces } of foraminiferal shell - they were slightly curved, crystalline-looking } shards with closely spaced fine openings (the foramen, for which forams are } named) and a few large holes (possible spine attachment localities). I ran } EDS on them to make sure that is indeed, what they were. Planktic foram } shells should be fairly pure calcium carbonate. What I got instead were } spectra each showing a fairly large strontium peak, a smaller sulfur peak } and a little oxygen. There were also very high aluminum peaks on each one, } but since the sample was simply air dried on an aluminum stub, I assume } that was the source of the Al. } In 20-some years of micropaleontology I've never heard of any marine } organism that uses strontium to build its shell. Sr is just below Ca in the } periodic table, so I suppose it shares some of the same reactability } characteristics (but I'm no chemist), so perhaps it could substitute, if no } Ca was available (which would be darn funny in a surface marine sample). } Let me add that all the other peaks were in their proper places, and the } instrument had been calibrated quite recently, and I can think of no source } of strontium contamination which could have gotten in there. We're planning } on sending some of these bits out for analysis by some other means just to } make sure, but for the time being, it's quite a mystery. } Any ideas from the marine biologists? The chemists? The EDS gurus? } Psychics? }
Matthew J. Lynn, Ph.D. Center for Advanced Microscopy University of Miami (305)284-4736 mlynn-at-miami.edu
Oldrich- I am sorry, but I have forgotten all of the details of your problem. Have you thermally cycled your detector? letting the detector be a room temperature for at least 24hrs can remove ice build-up in the detector. It can also be useful to purge the dewar with nitrogen at that time to remove any ice that may be in the dewar thus reducing cooling efficiency. I would also suggest cleaning your detector window (I don't know what kind you have.) Be very sure that you are using the proper technique and chemicals for cleaning the window as the polymer windows are extremely easy to damage (i.e., don't blow a dust particle off of it, see the current thread on particulate damage to EDX windows.) In my case, as my window gets coated with pump oil from the vacuum, etc. the Cu L line reduces in intensity relative to the K line. For my super thin window the ratio should be L:K } =2:1. If I put a thin graphite sheet between the detector and the sample, I get spectra very similar to the ones you have posted with the low energy x-rays blocked out (you'll still have the electronic noise peak at the lowest energy) and then a normal looking higher energy range. Perhaps some of this will help. I'll be on vacation the next couple of weeks, but give me a call after 7 July if you have any questions I can help with. Matthew Ervin MErvin-at-ARL.mil 301-394-0017
Oldrich Benada {benada-at-biomed.cas.cz} on 06/23/2000 05:11:15 AM
To: Microscopy-at-sparc5.microscopy.com cc:
Dear colleagues, Many thanks for all responses to my question considering the hole in the EDX spectrum. We had a service engineer in our lab and he tried to solve the problem but he was not successful. I have prepared WEB page with the images of spectra, that were recorded from following samples: Cu_Al sample (usually used for calibration) Ti grid AL grid K standard
The url of that page is following: http://www.biomed.cas.cz/~benada/EDX_page/index.htm
Please, if anyone would be so kind and could comment the spectra, we will be very happy. Best regards from Prague Oldrich Benada
+-----------------------------------+ Oldrich Benada Acad. Sci. CR Institute of Microbiology Laboratory of electron microscopy Videnska 1083 CZ - 142 20 Prague 4 - Krc Czech Republic +------------------------------------+ Phone: +420-2-4752399 Fax: +420-2-4715743 WEB:http://www.biomed.cas.cz/mbu/lem113/lem.htm
To all: We have been using Kodabrome II RC paper F1-F4 for years with our Durst Laborator 1200 and Rapidoprint DD3700. As of the end of 1999, the paper is still available and gives good results.
Peggy Sherwood Photopathology Wellman Labs of Photomedicine-W224 70 Blossom Street Boston, MA 02114
Does anyone know how a really good counterfeiter would make a change in the date field of a rare coin look as though it weren't altered? I've looked for markings associated with moving the metal around, a joint between the numbers and the base coin, and differing metallurgy, with no "smoking gun"....
Any thoughts??
Brian ---------------------------------------------- Brian McIntyre Electron Microscopy Lab- River Campus Univ of Rochester Rochester, NY 14627 716-275-3058/4875
Sorry if this is double-posted....server troubles!
Frank,
Yes...very interesting result! My guess is that you have happened upon a very rare group of microplankton which make a celestite test (SrSO4). If I remember my micropaleo, look for a group called Acantharia (?) Try Haq & Boersma's "Marine Micropaleontology" for a start. The reason you have not come across them in 20-odd years is that the celestite tests are very unstable...I saw a few beautiful examples in filtered water from the Arabian Sea, never in sediment samples. So not much use for paleoclimatology, but an interesting diversion. If you can post some pics to a website, I'd like to take a look at them.
Matt
On Friday, June 23, 2000 6:59 AM, Frank Thomas [SMTP:thomasf-at-AGC.BIO.NS.CA] 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 } -----------------------------------------------------------------------. } } } Here's an interesting problem. The other day I was examining some plankton } trap samples in our ESEM, and found a few bits of what I took to be pieces } of foraminiferal shell - they were slightly curved, crystalline-looking } shards with closely spaced fine openings (the foramen, for which forams are } named) and a few large holes (possible spine attachment localities). I ran } EDS on them to make sure that is indeed, what they were. Planktic foram } shells should be fairly pure calcium carbonate. What I got instead were } spectra each showing a fairly large strontium peak, a smaller sulfur peak } and a little oxygen. There were also very high aluminum peaks on each one, } but since the sample was simply air dried on an aluminum stub, I assume } that was the source of the Al. } In 20-some years of micropaleontology I've never heard of any marine } organism that uses strontium to build its shell. Sr is just below Ca in the } periodic table, so I suppose it shares some of the same reactability } characteristics (but I'm no chemist), so perhaps it could substitute, if no } Ca was available (which would be darn funny in a surface marine sample). } Let me add that all the other peaks were in their proper places, and the } instrument had been calibrated quite recently, and I can think of no source } of strontium contamination which could have gotten in there. We're planning } on sending some of these bits out for analysis by some other means just to } make sure, but for the time being, it's quite a mystery. } Any ideas from the marine biologists? The chemists? The EDS gurus? } Psychics? }
Matthew J. Lynn, Ph.D. Center for Advanced Microscopy University of Miami (305)284-4736 mlynn-at-miami.edu
A really good counterfeiter could do this at the molecular level by using and FIB or equivalent.
I haven't really explored exactly how this could be done but it has crossed my mind.
regards,
Earl Weltmer
Brian McIntyre 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 all- } } Does anyone know how a really good counterfeiter would make a change in the } date field of a rare coin look as though it weren't altered? I've looked } for markings associated with moving the metal around, a joint between the } numbers and the base coin, and differing metallurgy, with no "smoking gun".... } } Any thoughts?? } } Brian } ---------------------------------------------- } Brian McIntyre } Electron Microscopy Lab- River Campus } Univ of Rochester } Rochester, NY 14627 } 716-275-3058/4875
There was a recent posting about UA precipitating during staining.
Well wouldn't you know I too have experinecing similar problem. It took me a long time to figure out but I narrowed it down to our rinsing water that is used to rinse the grid in after it is stained.
After chaging the filters on our water purification system the problem was gone.
Rajesh Patel Robert Wood Johnson Medical School Dept. of Pathology Electron Microscopy Lab 675 Hoes Lane Piscataway, NJ 08854
how about looking at sub-surface damage (using some channeling techniques or similar)? I could imagine, that the damage from stamping the coin is different from Ion bombarding the coin.
Or, look for implanted atoms. The FIB process uses metal atoms to bombard the sample. Some of them get implanted into the material. So you might find higher concentrations where the sample was altered.
Finally, how about a surface layer like an oxide or patina. There may be differences between the very old layer and the new layer.
Michael
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 =================================== phone: (888) FIND SIS (303) 234-9270 fax: (303) 234-9271 email: mailto:info-at-soft-imaging.com web: http://www.soft-imaging.com ===================================
-----Original Message----- } From: Brian McIntyre [mailto:mcintyre-at-optics.rochester.edu] Sent: Friday, June 23, 2000 12:14 PM To: Microscopy-at-sparc5.microscopy.com
Hi all-
Does anyone know how a really good counterfeiter would make a change in the date field of a rare coin look as though it weren't altered? I've looked for markings associated with moving the metal around, a joint between the numbers and the base coin, and differing metallurgy, with no "smoking gun"....
Any thoughts??
Brian ---------------------------------------------- Brian McIntyre Electron Microscopy Lab- River Campus Univ of Rochester Rochester, NY 14627 716-275-3058/4875
Over time I saw some questions regarding the installation of a UPS or some power conditioner to improve the stability of all sorts of microscopy equipment. I understand that quite a few instruments are sensitive to the quality of the line power they get. As I was working on EMC (electro magnetic compatibility) in a former life, this seems strange to me. In Europe we have very strict EC-rules that guarantee that equipment meets its specifications even under severe conditions (including HF noise on the power, fluctuations in the power, high EM fields, temperature variations etc)
I agree that microscopes are far more sensitive devices than, say, videogames but does anyone know of the EMC rules for microscopes and related equipment? It seems strange that some people are using UPS's while especially power fluctuations can be easily overcome with a good design of the electronics...
Regards,
Jo
************************************************************* * Jo Verbeeck * * University of Antwerp * * Dept. EMAT (Electron Microscopy for Materials Research) * * e-mail: joverbee-at-ruca.ua.ac.be * * tel: +32(0)3 218 02 49 * * fax: +32(0)3 218 02 57 * *************************************************************
I've got my references now: The protists with SrSO4 tests are the class *Acantherea*, members of the phylum Sarcodina (amoebae), subphylum Actinopoda (used to be known as the Radiolaria).
By my somewhat outdated reference ("A Synoptic Classification of Living Organisms", R.S.K. Barnes, ed.). The names have likely been changed, and the Actinopoda broken up to protect the careers of systematists.
Forget I ever mentioned Actinaria or Acanthella. The long-term storage in my wetware is getting flakey (I misremembered).
Phil
}}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{ Philip Oshel Supervisor, AMFSC and BBPIC Dept. of Animal Health and Biomedical Sciences University of Wisconsin 1656 Linden Drive Madison, WI 53706-1581 Voice: (608) 263-4162 peoshel-at-facstaff.wisc.edu fax: (608) 262-7420 (dept. fax)
Could anyone recommend an independent U.K based company specialising in the repair and maintenance of electron microscopes ?
Without mentioning names I feel that the service we receive re our SEM maintenance contract is particularly poor and am beginning to wonder is there any alternative to complaint without improvement and excuse after excuse .
I have noticed that little repair is carried out at component level during service engineer visits but am not aware of how ' specific ' most parts are to a particular make of instrument . If parts always have to be ordered through the instrument supplier there would be no advantage in independent maintenance . Another alternative perhaps would be in-house maintenance but do courses exist to train the novice type like myself to a reasonably competent level or is this not advised ?
Keen to hear comments ...
M.Harris Email harrism-at-esm-semi.co.uk ESM LTD , South Wales , U.K
} Date: Mon, 26 Jun 2000 09:56:43 +0200 } To: Microscopy-at-sparc5.microscopy.com } From: Rudi Lurz {Lurz-at-molgen.mpg.de} } Subject: Re: ***TEM Darkroom Users*** AGFA Paper has become extinct ! } } To all: } The identical technology (developer in emulsion + activator) as with } Rapidoprint papers is used with Agfa Brovira-Speed papers. I have replaced } years ago the Rapitone paper by Brovira-Speed 310 RC because I prefered } more the blue-black of Brovira-Speed to Rapitone which showed more brown } tendency. Brovira-Speed is available from soft to extra hard and should be } still produced (hopefully) as I was told from my dealer this morning. } This is for Germany - I do not know the situation in the US. } Agfa seems to reduce drastically the B/W programme. The Agfa Scientia } 23D56 films are no longer produced too and we have to switch to Kodak SO-163. } Best regards } Rudi Lurz
_________________________________ Rudi Lurz Phone: X - 30-8413-1271 MPI fźr Molekulare Genetik Fax: X - 30-8413-1385 Ihnestrasse 73 D-14195 Berlin E-mail to: Lurz-at-molgen.mpg.de
Teaching microscopy at both the undergrad and grad student levels would be an excellent symposium or discussion group at the Long Beach meeting. Especially since the life of EM facilities depends on having users, and getting students interested in microscopy is one of the best ways to create users.
Phil
} } Greetings: } } I am a new subscriber to this list and am looking for input/advice } } regarding the use of SEM's in undergraduate research. I teach at a liberal } } arts university with only undergraduate programs in the natural sciences. } } A geology colleague and myself (I'm a biologist) are exploring possible } } funding sources which would allow us to purchase an SEM that would be used } } by biology and geology undergrads in their independent and senior thesis } } projects. (We currently have a non-functioning ETEC that we would like to } } replace -- we've been advised that further repair and head-banging would } } not be cost effective). If anyone is currently engaged in SEM-based } } research with undergrads, I would like to get your feedback on your } } experiences (both the pluses and minuses). Although we don't have a TEM, } } input from those using TEM with undergrads would also be helpful. } } } } Specific Questions: } } } } 1) What types of projects are your students pursuing? } } } } 2) Are you conducting interdisciplinary projects? e.g., micropaleontology } } or environmental applications. } } } } 3) Are there particular models of SEMs which would be more user friendly } } since we would be training undergrads and we would also be the primary } } trainers and technicians? } } } } 4) Have you developed an undergrad EM course? } } } } Thanks, } } Ken -- }}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{ Philip Oshel Supervisor, AMFSC and BBPIC Dept. of Animal Health and Biomedical Sciences University of Wisconsin 1656 Linden Drive Madison, WI 53706-1581 voice: (608) 263-4162 fax: (608) 262-7420 (dept. fax)
i should clarify that the coin in question could not have been altered after 1900 so modern techniques are obviously out of the question. the issue becomes one of... if you wanted to alter a coin's date how would you have done it 100 years ago, and how could it be uncovered using modern analysis techniques.
thanks! b- ---------------------------------------------- Brian McIntyre Electron Microscopy Lab- River Campus Univ of Rochester Rochester, NY 14627 716-275-3058/4875
Use of an x-ray microprobe should be rather definitive, I would think. While there are exotic (and expensive) techniques that could be used to remove a raised date and place a new one, it would be extremely difficult to exactly match the elemental contents, particlarly those of the trace elements. For example, gold, even when refined to 99.999% pure, contains enough impurities for any batch to be matched to a particular mine that it came from.
The only other method I could imagine is a close characterization of the grain structures. I don't know if the stamping process would have much of an affect on the surface grain, but I imagine any implantation method would leave a much different structure as completely different conditions were used to form the metal.
Allen R. Sampson, Owner Advanced Research Systems, St. Charles, Illinois 60174 voice 630.513.7093 fax 630.513.7092
On Friday, June 23, 2000 1:14 PM, Brian McIntyre [SMTP:mcintyre-at-optics.rochester.edu] wrote:
} Hi all- } } Does anyone know how a really good counterfeiter would make a change in the } date field of a rare coin look as though it weren't altered? I've looked } for markings associated with moving the metal around, a joint between the } numbers and the base coin, and differing metallurgy, with no "smoking gun".... } } Any thoughts?? } } Brian } ---------------------------------------------- } Brian McIntyre } Electron Microscopy Lab- River Campus } Univ of Rochester } Rochester, NY 14627 } 716-275-3058/4875 } }
A couple of weeks ago I wrote in witha problem trying to fix plant vesicle preps for LR White inclusion and immunogold studies, without osmicating. The pellets were disappearing during dehydration. I included an osmium step (5, 30 or 60 minutes) and the pellets stopped disappearing, so I guess there is so little protein that a formaldehyde/gluteraldehyde fix doesn't manage to stabilize them. I'll have to check whether these osmicated vesicles work in the immunogold reaction and check the effect of a metaperiodate treatment to expose the proteins. Thanks for the advice - another problem solved.
Someone brought us some frogs' eggs to fix. It looks like they have a hard skin. The researcher is interested in extracting the nuclei (huge) which also look like they're enclosed in a hard skin, and he wants to know whether the extracted nuclei still retain some endoplasmic reticulum, so I thought the best way would be to compare a whole egg with an extracted nucleus. Any ideas/experience on how to fix this thing?
Thanks,
Mark
******************************************** Mark West, Unidad de Microscopia Electronica, (Electron Microscopy Unit) Instituto de Fisiologia Celular, Universidad Nacional Autonoma de Mexico, 04510 Mexico D.F.
I just want some opinions about the problem I got recently with our SEM.
We use this machine for electron beam lithography. This application is a little more sensitive than imaging. The problem I have now is, when I image at very low magnification, I can see the circular rim of the final section of the column. But from a couple of weeks, it changed suddenly, now the circular image is distorted like an oval shape and at second saturation point, normally only one position of gun alignment gives the maximam beam current, but I see there are clearly two distinct, well separated positions. We think something is blocking in the middle of the beam path, but want to know whether there is any other one who experienced similar thing and know the reason. I appreciate if anyone can give me his/her similar experince.
Coins have been faked outright and updated using modern dies, and assembled from two separate halves using solder. Coins whose value depends on date have been "enriched" by removing the original date and soldering in its place numbers of a more valuable date. This latter case might be discerned using scanning electron microscopy to examine surface texture and degradation and adjunct energy-dispersive analysis to detect the presence of solder.
James Martin Principal/Research Scientist Orion Analytical, LLC P.O. Box 550 Williamstown, MA 01267 www.orionanalytical.com
} i should clarify that the coin in question could not have been altered } after 1900 so modern techniques are obviously out of the question. the } issue becomes one of... if you wanted to alter a coin's date how would you } have done it 100 years ago, and how could it be uncovered using modern } analysis techniques. } } thanks! } b- } ---------------------------------------------- } Brian McIntyre } Electron Microscopy Lab- River Campus } Univ of Rochester } Rochester, NY 14627 } 716-275-3058/4875 }
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You don't have to wait that long! Go to the Philadelphia meeting web page search engine http://www.msa.microscopy.com/cgi-bin/M&M00Program.pl and enter "teaching microscopy". Steve Barlow has organized an excellent all-day session.
Caroline
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
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Clearly, I have several vested interests here! As I am answering this from home, I don't actually know as I write if you are specifically concerned about a JEOL SEM, although I will check when I get to the office tomorrow.
First, I would suggest you complain, with specifics, to the service manager of the relevant company. If that is ineffective, direct your complaint to the Salesperson covering your area - sales people, perhaps, have a much stronger vested interest in happy customers. A happy customer is possibly going to buy again in the future, an unhappy customer makes their feelings known to others and can influence future sales.
I hope you are not a JEOL user - if you are, please contact me directly and I will attempt to resolve your problems.
If the SEM in question is an older instrument, independent service companies will do a good job on all of the more routine problems. You should also be able to resolve these yourself, in principle, with a reasonable knowledge of vacuum technology and electronics.
I don't know where you might get training on SEM service - possibly Protrain?
The two main independent service companies that I am aware of operating in the UK are ISS and EOS, both based in the Manchester area but with engineers throughout most of the UK. Contact details from sources on the web and Microscopy & Analysis. If you contect me directly, I will forward you details.
Problems arise with newer instruments, especially PC-controlled ones. Self-service and independent service will cover the basics but without detailed training, circuit diagrams, software tools, it gets very difficult to handle anything more than the basics.
Incidentally, this is one of the disadvantages for customers of computer controlled EMs - it makes it easier for ALL manufacturers to restrict service to approved service engineers. Not that this happens just with scientific instruments - a similar issue exists with cars/automobiles. I believe there is/has been a court case in the US by independent auto-repair companies against the manufacturers to force them to release service software.
Even on older instruments, major problems can be difficult to resolve for anybody other than the manufacturer.
You will probably find that service via independents is cheaper than via the manufacturers. Specialist parts will have to be purchased from the manufacturer but even then, the overall cost will probably be lower. However, I would add that I don't believe any EM manufacturer is interested in providing anything other than value-for-money service and, to a great extent, you do get what you pay for - at least, you should! If you are paying less, overall, the service you get is probably less - it may not be obvious, and you may never see the difference but, in the long run, you will get what you pay for.
You also need to keep in mind that the more comprehensive service contracts provided by manufactuers contain a significant 'insurance' element. Major components are expensive - with a comprehensive service contract from the maufacturer, you will never know how expensive but with service via an independent, you may find out!
Hope that is useful.
Best regards, -- Larry Stoter JEOL (UK) Ltd Silver Court, Watchmead, Welwyn Garden City, AL7 1LT, United Kingdom tel: +44-(0)1707-377117, fax: +44-(0)1707-373254, e-mail: larrys-at-jeoleuro.com
It would very much help to know what sort of an SEM you are using and at what kV you are operating. I suspect you may have had a minor vacuum accident which caused one of your "spray apertures" to be blown out of position by inrushing air. Pull the liner tube (if your instrument has one) and reposition all the apertures. It might not be a bad idea to do a good cleaning job while inside the column.
Good luck.
Alex Greene SCIENTIFIC INSTRUMENTATION SERVICES, INC. PMB-499, 1807 West Slaughter Lane, Number 200 Austin, Texas 78748-6200 Phone: 512/282-5507 Fax: 512/280-0702
QUALITY ELECTRON MICROSCOPE REPAIR -----Original Message----- } From: Heejun Jeong {" hjjeong"-at-physics.purdue.edu} {Heejun Jeong {" hjjeong"-at-physics.purdue.edu} } To: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com}
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There are essentially two types of UPS systems. One acts in a passive mode and switches to battery backup when power fails. The other is an active mode which converts AC input to DC and uses this to generate stable AC output.
Power conditioners are typically microprocessor-controlled units which adjust taps on a transformer to maintain reasonably stable output voltage. These can also include traps for spikes and overvoltage conditions (Topaz for example).
The issue here is the stability of the basic potentials and currents which are fed to or are delivered to the electron gun. In this regard, I think that we are talking about very narrow margins of input voltage variation and output voltages. I think that one will have line sag no matter what--on occasion. Thus, the active UPS units bypass these conditions and provide a constant, stable AC supply to the SEM. Line conditioners are rated at +/- 6% down to +/- 3%. Active UPS can do better.... but their surge current capabilities are less than those of the conditioner units.
Consequently, there is not one, singularly perfect solution. But there are options which the user should consider.
gg
At 12:29 AM 6/26/00, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} From a practical point of view my experience with computer controlled UPS systems is that cause more problems than they solve. They provide a computer with good enough power in most cases. This power is not good enough for equipment that is not designed for considerable voltage changes and micro power losses.
Most of my experience comes from computer back up power supplies and a project that was measuring sunlight. Since it was sunlight and we needed fast response we did not have any low pass filter. When working on it in the lab I used filtered DC since I was having problems with the 120 Hz flicker of incandescent bulbs. I could see sags and glitches in the AC power through the UPS systems even with the filters. It was good enough for what I was doing but I could see a good deal difference in the stability of the UPS and a battery.
I have worked with a few radio systems that use AC to charge a battery bank and then power the radios from the batteries. These are far more reliable than UPS systems.
Gordon Gordon Couger gcouger-at-couger.com Stillwater, OK www.couger.com/gcouger 405 624-2855 GMT -6:00
} From: "Dr. Gary Gaugler" {gary-at-gaugler.com} } } There are essentially two types of UPS systems. One acts in } a passive mode and switches to battery backup when power } fails. The other is an active mode which converts AC input } to DC and uses this to generate stable AC output. } } Power conditioners are typically microprocessor-controlled units which } adjust taps on a transformer to maintain reasonably stable } output voltage. These can also include traps for spikes and } overvoltage conditions (Topaz for example). } } The issue here is the stability of the basic potentials and currents } which are fed to or are delivered to the electron gun. In this } regard, I think that we are talking about very narrow margins of } input voltage variation and output voltages. I think that one will } have line sag no matter what--on occasion. Thus, the active } UPS units bypass these conditions and provide a constant, } stable AC supply to the SEM. Line conditioners are rated at } +/- 6% down to +/- 3%. Active UPS can do better.... but their } surge current capabilities are less than those of the conditioner units. } } Consequently, there is not one, singularly perfect solution. But there are } options which the user should consider. } } gg }
} } } } Hello all, } } } } Over time I saw some questions regarding the installation of a UPS or some } } power conditioner to improve the stability of all sorts of microscopy } } equipment. } } I understand that quite a few instruments are sensitive to the quality of } } the line power they get. } } As I was working on EMC (electro magnetic compatibility) in a former life, } } this seems strange to me. } } In Europe we have very strict EC-rules that guarantee that equipment meets } } its specifications even under severe conditions (including HF noise on the } } power, fluctuations in the power, high EM fields, temperature variations } } etc) } } } } I agree that microscopes are far more sensitive devices than, say, } } videogames but does anyone know of the EMC rules for microscopes and } } related equipment? } } It seems strange that some people are using UPS's while especially power } } fluctuations can be easily overcome with a good design of the } } electronics... } } } } Regards, } } } } Jo
I am trying to locate a copy of a graph that shows the thicknesses of (glass) windows that can support atmospheric pressure against a vacuum for various radii of O ring support. Kaufman glass has one for high pressures for several glass types but it doesn't cover the range of atmospheric pressure and relatively small radii(1 to 3 cm). Pyrex, leaded glass, and quartz are the three principal materials of interest. Appreciate any leads.
Don Marshall
Donald J. Marshall Relion Industries P.O. Box 12 Bedford, MA 01730 Ph: 781-275-4695 FAX: 781-271-0252 email dmrelion-at-world.std.com
Cathodoluminescence, mass spectroscopy, electron beam technology
"A weed is a flower out of place."
(Please note: Do not send email with attachments to this address. Instead, send it to donbarlen-at-aol.com. Thank you.)
You may want to contact David Gard, Ph.D., Biology Department, University of Utah, Salt Lake City, UT. He has done a great deal of work with frog eggs.
good luck
Ramin Rahbari Pfizer Global Research & Development Worldwide Preclinical Safety 2800 Plymouth Road Ann Arbor, MI 48105 Voice (734) 622-3383 Fax (734) 622-5001 Ramin.Rahbari-at-WL.COM
-----Original Message----- } From: Mark West [mailto:mwest-at-ifcsun1.ifisiol.unam.mx] Sent: Monday, June 26, 2000 3:37 PM To: Microscopy-at-sparc5.microscopy.com
Hi,
A couple of weeks ago I wrote in witha problem trying to fix plant vesicle preps for LR White inclusion and immunogold studies, without osmicating. The pellets were disappearing during dehydration. I included an osmium step (5, 30 or 60 minutes) and the pellets stopped disappearing, so I guess there is so little protein that a formaldehyde/gluteraldehyde fix doesn't manage to stabilize them. I'll have to check whether these osmicated vesicles work in the immunogold reaction and check the effect of a metaperiodate treatment to expose the proteins. Thanks for the advice - another problem solved.
Someone brought us some frogs' eggs to fix. It looks like they have a hard skin. The researcher is interested in extracting the nuclei (huge) which also look like they're enclosed in a hard skin, and he wants to know whether the extracted nuclei still retain some endoplasmic reticulum, so I thought the best way would be to compare a whole egg with an extracted nucleus. Any ideas/experience on how to fix this thing?
Thanks,
Mark
******************************************** Mark West, Unidad de Microscopia Electronica, (Electron Microscopy Unit) Instituto de Fisiologia Celular, Universidad Nacional Autonoma de Mexico, 04510 Mexico D.F.
We're in the process of acquiring an EDS system for our JEOL 5600 and I'm curious about the experience of others with EDS systems on scopes without specimen airlocks (column vents to exchange specimens). Any thoughts/recommendations? I'm particularly concerned with contamination,
window integrity and the like. Are there any advantages/disadvantages of
Be vs. thin window detectors here?
Thanks,
Jim Ehrman
--
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
I've received a query from a climatologist who wants to photograph fine varves and/or laminations (~0.5 mm- 1 mm each layer) in carbonate (calcite) rich silt (or marl) through a binocular light microscope. The samples are flat, finely cut or polished sections that can be etched if necessary, but he's really looking for a staining method that would enhance the difference between the carbonate-rich layers and the clay-rich layers to make them more usefully photogenic. Can anyone advise us on how to proceed to best advantage?
Many thanks, Dee
*************************************************************** Dee Breger Mgr. SEM/EDX Facility Lamont-Doherty Earth Observatory 61 Route 9W Palisades, NY 10964 USA T: 914/365-8640 F: 914/365-8155
http://www.ldeo.columbia.edu/micro http://www.discovery.com/area/science/micro/micro1.html http://www.lsc.org/antarctica/front.html Journeys in Microspace (Columbia University Press, 1995)
We have two different models of thin window detectors on two different scopes. Our Hitachi 2460N does not have an airlock and is vented for every sample change. Our JEOL 840A has an airlock, but does get the chamber vented fairly often. We have only had one pin-hole develop in the detector on our 840 in the 15-20 instrument-years that we have had the detectors.
Therefore, I don't think you would be gaining much, if anything, by switching to a Be window. And you would be giving up that wonderful light element capability. I would be hard pressed not to be able to detect C and O. I would go for the thin window and be reasonably careful.
Warren S.
At 10:59 AM 6/27/2000 -0300, you wrote:
} Hi all, } } We're in the process of acquiring an EDS system for our JEOL 5600 and } I'm curious about the experience of others with EDS systems on scopes } without specimen airlocks (column vents to exchange specimens). Any } thoughts/recommendations? I'm particularly concerned with contamination, } } window integrity and the like. Are there any advantages/disadvantages of } } Be vs. thin window detectors here? } } Thanks, } } Jim Ehrman } } -- } } James M. Ehrman } Digital Microscopy Facility } Mount Allison University } Sackville, NB E4L 1G7 } CANADA } } phone: 506-364-2519 } fax: 506-364-2505 } email: jehrman-at-mta.ca } www: http://www.mta.ca/~jehrman }
} Could anyone recommend an independent U.K based company } specialising in the repair and maintenance of electron microscopes ? } } Keen to hear comments ...
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An investigator just brought me 6 samples - heads of flies. He was instructed to bring the entire body but he cuts the heads off when he does TEM. The investigator knows nothing about SEM. I have done alot of scanning of fly eyes but the body is attached. My question is: how do I mount the heads on the stubs without the graphite or silver paint causing capillary action over the entire head? We will scan on a JEOL 840A after Au/Pd coating.
George Lawton Chief Electron Microscopist Molecular and Cellular Imaging Facility UT Southwestern Medical Center at Dallas 5323 Harry Hines Blvd. Dallas, Tx 75390-9039 Phone: 214-648-7291 Fax 214-648-6408 eMail: George.Lawton-at-email.swmed.edu
I would use one of the double-sided carbon sticky tabs available from several vendors. They are good for small samples that would be buried in liquids. If they don't provide enough contact area, you could very carefully dab a conductive paint around the base of the mounted heads with a pointy stick.
Cheers, Randy
Randy Tindall EM Specialist Electron Microscopy Core Facility W122 Veterinary Medicine University of Missouri Columbia, MO 65211 Tel: (573) 882-8304 Fax: (573) 884-5414 Email: tindallr-at-missouri.edu Web: http://www.biotech.missouri.edu/emc/
-----Original Message----- } From: George Lawton [mailto:George.Lawton-at-email.swmed.edu] Sent: Tuesday, June 27, 2000 10:03 AM To: microscopy-at-sparc5.microscopy.com
An investigator just brought me 6 samples - heads of flies. He was instructed to bring the entire body but he cuts the heads off when he does TEM. The investigator knows nothing about SEM. I have done alot of scanning of fly eyes but the body is attached. My question is: how do I mount the heads on the stubs without the graphite or silver paint causing capillary action over the entire head? We will scan on a JEOL 840A after Au/Pd coating.
George Lawton Chief Electron Microscopist Molecular and Cellular Imaging Facility UT Southwestern Medical Center at Dallas 5323 Harry Hines Blvd. Dallas, Tx 75390-9039 Phone: 214-648-7291 Fax 214-648-6408 eMail: George.Lawton-at-email.swmed.edu
I have dug through the archives of this fine list and found a bunch of information on vibration and microscopy. I work for an engineering consulting company, and part of our work is related to vibration of various sorts (implosions, blasting, construction, pile driving, vehicular vibration). A current project is for a medical facility that is expanding, including operating rooms that have surgical microscopes. The facility is very close to a freight line, and we are working with them to provide mitigation measures before the facility is constructed.
My question to the list is, does anybody know of actual vibration criteria in terms of amplitude (peak or RMS) and frequency for ANY microscopes. The microscope manufacturers we have contacted indicate no research in the area, which is surprising to me.
Our previous work with both human perception and machine sensitivity has usually involved some specific vibration criterion, whether it be for damage, annoyance, or whatever purpose. The discussion of the various isolation tables on the list is interesting, including the innertubes and tennis balls. They appear to be a "One isolator fits all" approach, which, if it works, is also surprising to me. Any help or leads in this regard would be greatly appreciated. Thanks
Doug Anderson
Douglas A. Anderson, PhD Senior Consultant Schnabel Engineering Associates (http://www.schnabel-eng.com) 510 East Gay Street West Chester, PA 19380 Phone: 610 696-6066, Fax: 610 696-7771
Brian, that's a very intriguing problem. What is the metal or alloy of the coin? What is the particular date you are dealing with? Could a counterfeiter have done something so simple as cutting out segments of a pre-existing numeral to alter the date? If so, perhaps channelling would reveal the coining deformation pattern below the missing segments.
Also, is it clear that the WHOLE COIN is not a counterfeit? Ordinarily, the effort and cost to create a die set for counterfeiting a whole coin would deter anyone from the effort, unless the coin is spectacularly valuable. On the other hand, if someone had a genuine coin (or a carefully modified one) to use as a pattern, it would not be too dreadfully difficult (even a century ago) to make a ceramic mold directly, or a metal die indirectly, from the pattern. Then you could produce "knock-offs" from that die. And your "real McCoy" could be undamaged. This would take a clever, patient, and very meticulous craftsman to pull it off, however.
==================================== Roy Arrowood, Associate Professor Metallurgical and Materials Engineering UTEP, El Paso, TX 79968-0520 (915)747-6934 NEW E-MAIL ADDRESS: arrowood-at-miners.utep.edu
This worked for amphipod maxilla, which are minute or smaller and setose, so it should work for fly heads: 1) mount on double-sticky carbon-conductive tabs; cut these from a sheet, or buy as stub-sized tabs 2) sharpen toothpicks or other sticks, some to a point and others to a pen nib (with or without the slot found in a fountain pen nib) 3) place a small blob of Ag paint at the edge of the stub connecting the surface of the sticky tab to the metal of the stub; I prefer Ag paint dissoved in methylethylketone 4) use the stick to draw the paint from the blob to the fly head (etc.); connect this to a part of the head that you don't care about; draw a circle around the head just far enough not to touch the head -- this creates a shorter conductive path from the specimen to the Ag paint
Careful use of the sticks and paint will allow you to use Ag paint on very small, hairy specimens. The paint can be allowed to partially dry (solvent to evaporate) to thicken it, but this may just form a skin on the surface, and does lead to stringing of the partly dry paint which can be very annoying.
Phil
} An investigator just brought me 6 samples - heads of flies. He was } instructed to bring the entire body but he cuts the heads off when } he does TEM. The investigator knows nothing about SEM. I have done } alot of scanning of fly eyes but the body is attached. } My question is: how do I mount the heads on the stubs without the } graphite or silver paint causing capillary action over the entire } head? We will scan on a JEOL 840A after Au/Pd coating. } } George Lawton } Chief Electron Microscopist } Molecular and Cellular Imaging Facility } UT Southwestern Medical Center at Dallas } 5323 Harry Hines Blvd. } Dallas, Tx 75390-9039 } Phone: 214-648-7291 } Fax 214-648-6408 } eMail: George.Lawton-at-email.swmed.edu
-- }}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{ Philip Oshel Supervisor, AMFSC and BBPIC Dept. of Animal Health and Biomedical Sciences University of Wisconsin 1656 Linden Drive Madison, WI 53706-1581 voice: (608) 263-4162 fax: (608) 262-7420 (dept. fax)
{bold} {bigger} {bigger} Microscopy and Molecular Biology Postdoctoral Position
Department of Zoology
Miami University
Oxford, OH {/bold} {FontFamily} {param} TIMES {/param}
{FontFamily} {param} Arial {/param} A two-year postdoctoral position is available for investigating the role of inner arm dyneins in ciliary motility. We have seven different inner arm dynein heavy chain genes cloned from {underline} Tetrahymena {/underline} {underline} thermophila {/underline} and are in the process of creating knockout mutations in those genes. The successful applicant will be involved in all aspects of the work, including creation of the knockout mutants and analysis of the phenotypes. The successful candidate should be experienced in light and electron microscopy and willing to learn molecular biology and some protein biochemistry (preferred) or experienced in molecular biology and protein biochemistry and willing to learn light and electron microscopy. Send letter, curriculum vitae, and reference letters to:
David Pennock
Department of Zoology
Miami University, Oxford, OH 45056
Email: pennocdg-at-muohio.edu {FontFamily} {param} TIMES {/param} {smaller} {smaller}
Drill small holes in stubs and glue a dress making pin in each hole spike upwards. Coat this in a sputter coater.
Push each insect head onto a pin and sputter coat at a specimen-target distance of 5cms.
This technique is used by a number of our clients.
Steve Chapman Senior Consultant Protrain For consultancy and professional training in EM world wide. www.emcourses.com Tel 44+ 1280 814774 Fax 44+ 1280 814007
I've done a little work with varves...let me start by saying that typically this imaging is done using X-rays or by making conventional thin sections for petrographic microscopy. That said, I vaguely recall staining carbonate thin sections with Alizarin Red S. There is a different stain for dolomite. I'm not sure how well the technique can be applied to large sections....are they still wet? Dehydrated? Vacuum impregnated? Let me know if this is on the right track and I will hunt for some references.
Matt
Matthew J. Lynn, Ph.D. Center for Advanced Microscopy University of Miami (305)284-4736 mlynn-at-miami.edu
On Tuesday, June 27, 2000 10:21 AM, Dee Breger [SMTP:micro-at-ldeo.columbia.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 colleagues, } } I've received a query from a climatologist who wants to photograph fine } varves and/or laminations (~0.5 mm- 1 mm each layer) in carbonate (calcite) } rich silt (or marl) through a binocular light microscope. The samples are } flat, finely cut or polished sections that can be etched if necessary, but } he's really looking for a staining method that would enhance the difference } between the carbonate-rich layers and the clay-rich layers to make them } more usefully photogenic. Can anyone advise us on how to proceed to best } advantage? } } Many thanks, } Dee } } } } } *************************************************************** } Dee Breger } Mgr. SEM/EDX Facility } Lamont-Doherty Earth Observatory } 61 Route 9W } Palisades, NY 10964 USA } T: 914/365-8640 } F: 914/365-8155 } } http://www.ldeo.columbia.edu/micro } http://www.discovery.com/area/science/micro/micro1.html } http://www.lsc.org/antarctica/front.html } Journeys in Microspace (Columbia University Press, 1995) }
Dear Jim, We have a Kevex Quantum light element EDX on a Hitachi that vents the whole chamber and it has been no problem, but I am told that the rate of venting can make quite a difference. I bought our EDX detector used and the previous owner claimed to have replaced the window every year for the five years that he had it. I have never had to replace the window and that is in a student lab. This particulars SEM takes over a minute to vent and makes no discernable sound, whereas my other SEM vents in ten seconds with an audible hiss. It, fortunately, has a Be-window detector, which is definitely tougher. If your SEM vents quickly, you might try to put some sort of restricter or filter on the vent to clean and slow it down. At 10:59 AM 6/27/00 -0300, you wrote: } } Hi all, } } We're in the process of acquiring an EDS system for our JEOL 5600 and } I'm curious about the experience of others with EDS systems on scopes } without specimen airlocks (column vents to exchange specimens). Any } thoughts/recommendations? I'm particularly concerned with contamination, } } window integrity and the like. Are there any advantages/disadvantages of } } Be vs. thin window detectors here? } } Thanks, } } Jim Ehrman Regards, Mary
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-interchg.ubc.ca
A colleague of mine has asked me to post the follwoing job announcement for an opening in our Cancer Research area. Please send resumes to the address at the bottom of the ad, or visit the website, which is also at the bottom of the ad. Please do not contact me directly concerning this position. Thanks!
Jane A. Fagerland, Ph.D. Dept. Microscopy and Microanalysis Abbott Laboratories
ABBOTT LABORATORIES PHARMACEUTICAL PRODUCTS DIVISION Abbott Park, Illinois
Abbott Laboratories is a global diversified company dedicated to the discovery, development, manufacture and marketing of health care products and services. Around the world, the company,s 54,000+ employees are committed to improving people,s lives by providing cost-effective health care technologies. We are interested in interviewing candidates for the following opportunity:
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Please send resumes to: 100 Abbott Park Road, D-583, AP9A, Abbott Park, Illinois 60064 Attn. Job Code 2K-KDA3422. Or visit our web site at www.abbott.com
Before staining, try using a UV light source to see if the calcite fluoresces - can fluoresce red to pink, orange, white, yellow or blue.
If you need to stain, use Alizarin Red S for staining. If it turns a deep red color, it's most likely calcite. Procedures for this can be found in "Laboratory Handbook of Petrographic Techniques" by Charles S. Hutchinson, 1974, John Wiley and Sons. Or you can have a thin section preparation lab do the staining for you - this will be much easier since they are set up to do this routinely.
Cheers, James Talbot
K/T GeoServices, Inc. X-ray diffraction petrologic studies visit my web site at http://www.ktgeo.com Argyle, TX, USA, (214) 403-6342
} ----- Original Message ----- } From: Dee Breger {micro-at-ldeo.columbia.edu} } To: {microscopy-at-sparc5.microscopy.com} } Sent: Tuesday, June 27, 2000 9:20 AM } Subject: LM geological stain question } } } } ------------------------------------------------------------------------ } } 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 colleagues, } } } } I've received a query from a climatologist who wants to photograph fine } } varves and/or laminations (~0.5 mm- 1 mm each layer) in carbonate } (calcite) } } rich silt (or marl) through a binocular light microscope. The samples are } } flat, finely cut or polished sections that can be etched if necessary, but } } he's really looking for a staining method that would enhance the } difference } } between the carbonate-rich layers and the clay-rich layers to make them } } more usefully photogenic. Can anyone advise us on how to proceed to best } } advantage? } } } } Many thanks, } } Dee } } } } } } } } } } *************************************************************** } } Dee Breger } } Mgr. SEM/EDX Facility } } Lamont-Doherty Earth Observatory } } 61 Route 9W } } Palisades, NY 10964 USA } } T: 914/365-8640 } } F: 914/365-8155 } } } } http://www.ldeo.columbia.edu/micro } } http://www.discovery.com/area/science/micro/micro1.html } } http://www.lsc.org/antarctica/front.html } } Journeys in Microspace (Columbia University Press, 1995) } } } } } } } } }
I've mounted small insects similar to what you now have using the following method:
1. Get some dissecting pins, preferably, or any very thin, rigid pin or wire and cut about 1/4 inch long.
2. Mount it vertically on an SEM stub using carbon paint, or any quick drying glue, and let it dry completely, put in warming oven to speed it up, or use hair dryer.
3. Position the fly heads upsidedown on clean surface, like lens tissue or glass petri dish.
4. Put a fresh drop of carbon paint, or glue, on a nearby surface, pick up the SEM stub with stub handling forceps, dip the tip of the pin into the fresh paint or glue just enough to get a tiny little blob on the pin head, then under a dissecting or low power scope touch the tip of the pin to the back surface of the fly head. It should stick and you can pick it up at that point, place right side up and let dry completely.
5. Coat in vacuum evaporator or sputter coater as usual. After coating run, vent gas into chamber slowly to not disturb delicate sample.
Its a bit of a prep, but you also get the fly head off the stub surface, little or no background junk in the view. Good luck!
Gib
Responding to the message of {s9587bef.020-at-mednet.swmed.edu} from "George Lawton" {George.Lawton-at-email.swmed.edu} : } } An investigator just brought me 6 samples - heads of flies. He was } instructed to bring the entire body but he cuts the heads off when he does } TEM. The investigator knows nothing about SEM. I have done alot of scanning } of fly eyes but the body is attached. } My question is: how do I mount the heads on the stubs without the graphite } or silver paint causing capillary action over the entire head? We will scan } on a JEOL 840A after Au/Pd coating. } } George Lawton } Chief Electron Microscopist } Molecular and Cellular Imaging Facility } UT Southwestern Medical Center at Dallas } 5323 Harry Hines Blvd. } Dallas, Tx 75390-9039 } Phone: 214-648-7291 } Fax 214-648-6408 } eMail: George.Lawton-at-email.swmed.edu } } } .
Gib Ahlstrand Electron Optical Facility, University of Minnesota, Dept. Plant Pathology 495 Borlaug Hall, St. Paul, MN. USA. 55108 (612)625-8249 612-625-9728 FAX, giba-at-puccini.cdl.umn.edu http://biosci.umn.edu/MIC/consortium.html
Just a reminder while we are on the subject of thin window detectors - as well as controlling the gas flow on venting also make sure that the chamber can not overpressure. On SEMs the chamber door should be able to open freely as soon as it reaches atmospheric pressure. On TEMs an aperture mechanism or similar should be freed so that it can release at atmospheric pressure.
Ron
On Tue, 27 Jun 2000, 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 Jim, } We have a Kevex Quantum light element EDX on a Hitachi that vents the whole } chamber and it has been no problem, but I am told that the rate of venting } can make quite a difference. I bought our EDX detector used and the previous } owner claimed to have replaced the window every year for the five years that } he had it. I have never had to replace the window and that is in a student } lab. This particulars SEM takes over a minute to vent and makes no } discernable sound, whereas my other SEM vents in ten seconds with an audible } hiss. It, fortunately, has a Be-window detector, which is definitely } tougher. If your SEM vents quickly, you might try to put some sort of } restricter or filter on the vent to clean and slow it down. } At 10:59 AM 6/27/00 -0300, you wrote: } } } } Hi all, } } } } We're in the process of acquiring an EDS system for our JEOL 5600 and } } I'm curious about the experience of others with EDS systems on scopes } } without specimen airlocks (column vents to exchange specimens). Any } } thoughts/recommendations? I'm particularly concerned with contamination, } } } } window integrity and the like. Are there any advantages/disadvantages of } } } } Be vs. thin window detectors here? } } } } Thanks, } } } } Jim Ehrman } Regards, } Mary } } 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-interchg.ubc.ca } } }
=========================================================================== Mr. Ron Doole e-mail ron.doole-at-materials.ox.ac.uk Department of Materials, phone +44 (0) 1865 273701 University of Oxford, fax +44 (0) 1865 283333 Parks Road. Oxford. OX1 3PH. UK. ============================================================================
OK thanks for the change of address, as usual we are looking for: books on microscopes catalogs of microscopes microscope instruction manuals books on application of microscopes either in biological or material sciences .. anything! Optical, em sem, afm, etc etc etc.
----- items and literature relating to any form of electrical communication and engineering of thereof (yes telephone, telegraph, early wireless telegraphy ---- artifacts and literature on radio and TV broadcasting --- Radar and radar countermeasures ----- cryptanalysis --------
thanks in advance as always Ed Sharpe archivist for SMECC
I have used my Umax Powerlook III flatbed scanner with transparency adapter many times to get high quality scans of my TEM negatives but today it is driving me crazy. I have some great gold labeling of pretty electron dense granules. On the negative I can clearly see the gold against the granule matrix but when I scan, the 256 grey levels compresses all that info into one level of black and therefore I can't discern the gold. any scan gurus out there who can help? I have tried playing with the gamma without much luck. thanks in advance, tom
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility
3 Tucker Hall Division of Biological Sciences University of Missouri Columbia, MO 65211-7400 (573)-882-4712 (voice) (573)-882-0123 (fax)
Tom, Have you tried telling the scanner that the image is a positive transparency rather than a negative? And then reverse the contrast in photoshop rather than with the scanner software. Greg
At 12:20 PM 06/28/2000 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Gregory W. Erdos, Ph.D. Assistant Director, Biotechnology Program PO Box 110580 University of Florida Gainesville, FL 32611
we have a Epson 1600, sometimes this problem happens too. my solution is to scan the negative in positive mode first then do a inversion after that. Hope this helps.
Hao Li
----- Original Message ----- } From: "Tom Phillips" {PhillipsT-at-missouri.edu} To: {Microscopy-at-sparc5.microscopy.com} Sent: Wednesday, June 28, 2000 1:20 PM
Hi, Thomas
I find it best to scan a negative in as a positive transparency at 6 million colors (rather than the 256 greys you are using), and then reverse it. This gives me the best tonal range.
Aloha, Tina
} } I have used my Umax Powerlook III flatbed scanner with transparency } } adapter many times to get high quality scans of my TEM negatives but } } today it is driving me crazy. I have some great gold labeling of } } pretty electron dense granules. On the negative I can clearly see } } the gold against the granule matrix but when I scan, the 256 grey } } levels compresses all that info into one level of black and therefore } } I can't discern the gold. any scan gurus out there who can help? I } } have tried playing with the gamma without much luck. thanks in } } advance, tom } }
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
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Oxford Instruments supply a wide range of CL detection systems, from low cost broad-band detectors to multiple-band, UV, IR and spectroscopic systems. As far as I am aware, they are the only company supplying such a wide range of CL systems - if there are others, sorry, and I would also like to know!
Regards -- Larry Stoter JEOL (UK) Ltd Silver Court, Watchmead, Welwyn Garden City, AL7 1LT, United Kingdom tel: +44-(0)1707-377117, fax: +44-(0)1707-373254, e-mail: larrys-at-jeoleuro.com
Thanks for all the suggestions. We did indeed find that scanning in the color mode (which is 14 bit) solved the problem. I don't know why UMax has their greyscale image only go to 256 colors. I will ultimately incorporate this in a lecture when i am teaching image analysis since it clearly shows the limitations of 8 bit images. once again, thanks.
tom
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility
3 Tucker Hall Division of Biological Sciences University of Missouri Columbia, MO 65211-7400 (573)-882-4712 (voice) (573)-882-0123 (fax)
Stephan, I know that Oxford instruments can supply you with the necessary equipment to do CL in the SEM. I am not sure whether this is standard equipment or whether it is special order, but I have seen it done. The equipment usually involves a parabolic mirror and a light-pipe with the relevant detector on the end of this. Then there is the software .etc. There is a contact email address on their web page:
http://www.oxford-instruments.com/
give them a try and see what they say.
Regards, Jonathan
******************************************************** Dr Jonathan Barnard
Analytical Materials Physics The Angstrom Laboratory, Uppsala University P O Box 534, SE-751 21 Uppsala, Sweden Phone: +46-(0)18-4716838 Fax: +46-(0)18-500131 Phone: Microscope room +46 18 471 6365 http://www.angstrom.uu.se/analytical/home.html ********************************************************
Has anyone tried Kodak's MDS 100 imaging system? I have a user who needs a quick documentation system and this might work for her. She is using NIH Image on a compound scope in our lab and she wants to become independent and do the work in her own lab, and maybe at other locations.
As an alternative, anyone with a Nubus frame grabber, ie Scion LG3 or equiv., that would sell it to her to use with NIH Image?
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
} I have used my Umax Powerlook III flatbed scanner with transparency } adapter many times to get high quality scans of my TEM negatives but } today it is driving me crazy. I have some great gold labeling of } pretty electron dense granules. On the negative I can clearly see } the gold against the granule matrix but when I scan, the 256 grey } levels compresses all that info into one level of black and therefore } I can't discern the gold. any scan gurus out there who can help? I } have tried playing with the gamma without much luck. thanks in } advance, tom } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } Tom -
You can get a really good book on scanner use at www.scantips.com or download it as a pdf,if you have the patience for 218 pp.
Caroline
Caroline Schooley Project MICRO Coordinator Microscopy Society of America Box 117, 45301 Caspar Point Road Caspar, CA 95420 Phone/FAX (707)964-9460 Project MICRO: http://www.msa.microscopy.com/ProjectMicro/PMHomePage.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/marinelab.html
Can anyone advise what the water flows should be in the three parallel branches (OL; DPs; and power boards) of the 840A) The manual spec is } 5 l/min total, but I need to know particularly what the flow should be thru the OL.
TIA
rtch
ps replies welcome from JEOL
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
We have an Oxford MiniCL detector attached to a JEOL 6300 - same column as the 6400. This is an inexpensive detector which suits our needs but there are other suppliers. No matter who you obtain one from, give some thought to what ports are free and to your analytical working distance (presume you have EDS or WDS). On the 6400 this is most likely 15mm - some ingenuity from Oxford or other supplier will be needed to clear the pole-piece eg an inclined configuration. It would be worth it - rather than have to alter the sample height for quant analysis.
Regards
Stafford Stafford McKnight Geology & Metallurgy University of Ballarat ph 03 53279262 fax 03 53279144
Accurel Systems International is looking for SEM Analysts: junior to experienced technicians and engineers
Responsibilities include all aspects of commercial SEM and EDX analysis (instrument operation, sample preparation, customer contact and follow-up, data interpretation etc.).
Knowledge of IC device structure, IC fabrication and packaging a plus.
2+ years hands on experience in the semiconductor industry preferred.
To find out more please check our web site: www.accurel.com
SEM candidates should send resume with references to Regina Campbell: reginac-at-accurel.com or FAX 408-737-3916.
Job description: You will form part of a team of specialists in analytical techniques, providing support to the rapidly growing III-V optoelectronic and microwave device fabrication plant at Caswell Technology. We perform a wide range of activities, including optical microscopy, SEM, TEM, EDX, X-ray techniques and scanning probe microscopies. You will be expected to be able to be competent in several of these techniques and have/develop a degree of expertise in one or more of them. Experience of semiconductor device fabrication techniques would be an advantage.
The post is available immediately.
More details of our company are available at http://www.caswelltechnology.com/
Please contact me if you are interested or would like any further information.
============================================================== Richard Beanland Caswell Technology, Caswell, Towcester, Northants NN12 8EQ
Don't broadcast change of address notices to the Listserver. If you want to change your subscription address, just send a unsubscribe message, followed by a subscribe message...
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================================================================== Nestor J. Zaluzec Materials Science Division Building 212 Argonne National Lab 9700 S. Cass Ave Argonne, Illinois 60439 USA Tel: 630-252-7901, Fax: 630-252-4798 Email: Zaluzec-at-aaem.amc.anl.gov ================================================================== TPMLab: http://tpm.amc.anl.gov MMSite: http://www.amc.anl.gov ==================================================================
The box said "This program requires Win 95/98/NT or better..." so I bought a G3 Mac
I run a microscopy unit which is completely digital. I have a problem with printing out images. I am planning to upgrade cameras to 10 or 12 bit megapixel units, but am already restricted by printers. Laser printers seem to deal with greyscales by halftoning at 100-150dpi which is unsatisfactory for small prints. Are there any true greyscale printers available, that manage better than 300dpi and better than 256 grey levels? Is there any advantage in having more than 256 grey levels? Sorry if this is a bit dim, but there are so many self styled experts who give conflicting opinions I end up totally confused. Thank you. Richard Black Nottingham, England richardblack-at-cwcom.net
This is my first message to the list... Caroline, I looked at Scan site and would like to try PDF format... but I can seem to locate the pathway to download? Any thoughts or pointers?
Thanks for the suggestion it looks like a great site. Tim Lyden, Ph.D. Research Scientist
Departments of Internal Medicine/Immunology and Physiology/Cell Biology Ohio State University Davis Medical Research Center 480 W 9th Ave. Columbus, Ohio 43210
I would like to purchase a "Tunneling Electron Microscope".
I have been unable to locate a supplier. If you know of one please let me know. Thanks! Alesia White Darling Secretary, Dr. Amnon Katz University of Alabama Aerospace Engineering and Mechanics Box 870280/205 Hardaway Hall Tuscaloosa, Alabama 35487-0280 ph: 205-348-8525 fax: 205-348-7240 or 205-348-2094
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I have found that the gold does not show up well when you scan the negative itself. So for gold to show up in the scanned image, I usually make one print and scan the print. This works quite well. I guess we will still have to work in darkrooms for quite sometime even though the digital imaging is becoming popular.
Good luck,
Soumitra
} } } I have used my Umax Powerlook III flatbed scanner with transparency } adapter many times to get high quality scans of my TEM negatives but } today it is driving me crazy. I have some great gold labeling of } pretty electron dense granules. On the negative I can clearly see } the gold against the granule matrix but when I scan, the 256 grey } levels compresses all that info into one level of black and therefore } I can't discern the gold. any scan gurus out there who can help? I } have tried playing with the gamma without much luck. thanks in } advance, tom } } Thomas E. Phillips, Ph.D. } Associate Professor of Biological Sciences } Director, Molecular Cytology Core Facility } } 3 Tucker Hall } Division of Biological Sciences } University of Missouri } Columbia, MO 65211-7400 } (573)-882-4712 (voice) } (573)-882-0123 (fax)
***************************************************************** Soumitra Ghoshroy Ph.D. Electron Microscopy Lab Box 3EML New Mexico State University Las Cruces, NM 88003 Tel: 505-646-3600 Fax: 505-646-5665 e-mail: ghoshroy-at-nmsu.edu http://confocal.nmsu.edu/eml
As I recall, the human eye is hard pressed to distinguish more than 6-bit grayscale (64 levels) unless the shades are right next to each other, which they might be in an image. Therefore, printing at 256 gray levels should be more than adequate.
The 10 or 12-bit cameras will still be handy in storing the extra information in your images. You will be able to do some gray scale manipulation to highlight the darker or lighter regions of your image. But when it comes to print time, you will be throwing out all but about 8 bits.
You rightly say that halftoning is a problem on small prints. You need a 16x16 dot cell to render 256 gray levels in halftone. For 1200 dpi printing, that means that you can achieve 75 pixels per inch (not very high). That means a 1024 pixel image would require 13.65 inches to show all that resolution. If the image is printed out smaller, then either grayscale or pixel resolution would be sacrificed. (Printing at 64 gray levels would require a 8x8 cell and could be done at 150 pixels per inch so that the image could fit into 6.8 inches.)
I don't have a dye sub printer here, but their specs would probably meet your needs. You would want to see how well they can render the grayscale. Since they do not halftone, the dpi spec would be the same as the pixel per inch spec. A 300 dpi printer should be able to render full resolution on a 1024 pixel image in 3.4 inches. But check out the grayscale performance to see if it is adequate.
Warren S.
At 07:58 AM 6/29/2000 -0500, you wrote:
} I run a microscopy unit which is completely digital. I have a problem } with printing out images. I am planning to upgrade cameras to 10 or 12 } bit megapixel units, but am already restricted by printers. Laser } printers seem to deal with greyscales by halftoning at 100-150dpi which } is unsatisfactory for small prints. Are there any true greyscale } printers available, that manage better than 300dpi and better than 256 } grey levels? Is there any advantage in having more than 256 grey } levels? Sorry if this is a bit dim, but there are so many self styled } experts who give conflicting opinions I end up totally confused. } Thank you. } Richard Black } Nottingham, England } richardblack-at-cwcom.net
I think this query was posted recently but I haven't been able to dredge up the replies in the list archives - too recent, maybe.
I'm looking for contact info for a colleague, for Balzers or Bal-tec. He inherited a B. carbon evaporator model CED 030 (also: 3U-G03-751/132). Can anyone help out with either contact info for the company, or a copy of the manual?? We will pay copy/ship charges.
Thanks, much.
Ann Hein Lehman Manager, EM Facility Trinity College Hartford CT 06106 v. 860-297-4289 f. 860-297-2538 e. ann.lehman-at-trincoll.edu
Hi Hai Li: Here is some references which claim MgAl2O4 is a good substrate for growing perovskites:
Miura, S., Yoshitake, T., Matsubara, S., Miyasaka, Y., Shohata, N. and Satoh, T., Epitaxial Y-Ba-Cu-O Films on Si with Intermediate Layer by RF Magnetron Sputtering, Appl. Phys. Lett. 53:1967-1969 (1988).
Wu, X.D., Inam, A., Hegde, M.S., Wilkens, B., Chang, C.C., Hwang, D.M., Nazar, L., Venkatesan, T., Miura, S., Matsubara, S., Miyasaka, Y. and Shohata, N., High Critical Currents in Epitaxial YBa2Cu3O7-x Thin Films on Silicon with Buffer Layers, Appl. Phys. Lett. 54:754-756 (1989).
My buddy, Darrell Schlom at Penn State tells me that using perovskite substrates is much better:
Y. Jia, M.A. Zurbuchen, S. Wozniak, A.H. Carim, D.G. Schlom, L-N. Zou, S. Briczinski, and Y. Liu, "Epitaxial Growth of Metastable Ba2RuO4 Films with the K2NiF4 Structure," Applied Physics Letters 74 (1999) 3830-3832.
Best regards, Mike Urbanik www.crystalguru.com
} } Subj: information about MgAl2O4
For a tungsten emitter, my JEOL operators' manual implies to raise the anode closer to the Wehnelt for low keV applications (less than 15keV). The manual does not mention this distance for the LaB6 emitter option. Any suggestions?
TIA and cheerios, =shAf= :o)
{} /\ {\/} /\ {\/} /\ {\/} /\ cogito, ergo zZOooOM /\ {\/} /\ {\/} /\ {\/} /\ {} Michael Shaffer, R.A. - mshaf-at-darkwing.uoregon.edu Geological Science's Electron Probe Facility - University of Oregon http://epmalab.uoregon.edu/
We also have an Oxford MiniCL detector (attached to our JEOL-8900), however the Research Instruments group of Oxford (CL and Cryo EM stuff) was just sold to Gatan, so you may need to contact Gatan to find out about the product. We are in the middle of some modifications to our unit (extension of the light pipe), and JEOL informed me of the transaction. It doesn't appear on the websites, however it was official as of 6-19-00. Hope the info helps. Sarah
Stafford McKnight 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 } -----------------------------------------------------------------------. } } Stephan, } } We have an Oxford MiniCL detector attached to a JEOL 6300 - } same column as the 6400. This is an inexpensive detector which } suits our needs but there are other suppliers. } No matter who you obtain one from, give some thought to what } ports are free and to your analytical working distance (presume you } have EDS or WDS). On the 6400 this is most likely 15mm - some } ingenuity from Oxford or other supplier will be needed to clear the } pole-piece eg an inclined configuration. It would be worth it - rather } than have to alter the sample height for quant analysis. } } Regards } } Stafford } Stafford McKnight } Geology & Metallurgy } University of Ballarat } ph 03 53279262 } fax 03 53279144
-- Sarah A.W. Lundberg Electron Microanalysis and Imaging Laboratory Department of Geoscience, UNLV 4505 S. Maryland Parkway Box 454010 Las Vegas, NV 89154-4010
Hellow Richard, HP Laserjet 2100TN is used in our laboratory and it has a very high stated resolution of 1200dpi. The printer gives excellent output at its highest setting, and lower. The level of detail I obtain from the printer is sufficient to show all the information from TEM images. However, I believe the final output from a photograph is still slightly better. Also, photographs have very nice glossy or matt finishes which I haven't seen on laser printouts.
\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\ Gordon Ante Vrdoljak Electron Microscope Lab ICQ 23243541 http://nature.berkeley.edu/~gvrdolja 26 Giannini Hall gvrdolja-at-nature.berkeley.edu UC Berkeley phone (510) 642-2085 Berkeley CA 94720-3330 fax (510) 643-6207 cell (510) 290-6793
On Thu, 29 Jun 2000, richard black 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 run a microscopy unit which is completely digital. I have a problem } with printing out images. I am planning to upgrade cameras to 10 or 12 } bit megapixel units, but am already restricted by printers. Laser } printers seem to deal with greyscales by halftoning at 100-150dpi which } is unsatisfactory for small prints. Are there any true greyscale } printers available, that manage better than 300dpi and better than 256 } grey levels? Is there any advantage in having more than 256 grey } levels? Sorry if this is a bit dim, but there are so many self styled } experts who give conflicting opinions I end up totally confused. } Thank you. } Richard Black } Nottingham, England } richardblack-at-cwcom.net } } } }
Balzers stuff is now handled by a company called Techno Trade. If you can't find them on the 'net or find their address, email me and I'll dig through my files and find their address for you.
Cheers :o)
Paul Grover Chief Microscopist and Bottle Washer Microvista Laboratory Lafayette, IN
My Amray says the same thing. As long as the gun's Whenelt end is the same distance for each emitter type, the spacing should be the same. I used 4mm {=5KV, 6mm {=12KV and 8mm for } 12KV.
gary g.
At 02:47 PM 6/29/00, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I'm selling a perhaps incomplete set of parts for a phase contrast setup for a B&L microscope. I don't have any B&L equipment, and it was part of a lot I got a sale....
ADI - FOUNDRYS OFFER FOR DESIGNERS AND USERS OF CASTINGS
23-24 NOVEMBER 2000 CRACOW – FOUNDRY RESEARCH INSTITUTE
CIRCULAR NO. 1
HOSTED BY: Instytut Odlewnictwa 30-418 Kraków, Zakopiańska 73
CO-ORGANISER: Ministry of Economy Polish Foundrymen’s Technical Association Foundry Economic Chamber SCIENTIFIC COMMITTEE
List of persons invited to participation in the Scientific Committee of the Conference: Prof. Jerzy PIASKOWSKI – Poland Prof. Czesław PODRZUCKI – Poland Prof. Edward GUZIK – Poland Prof. Jan RĄCZKA – Poland Prof. Ryszard KOZŁOWSKI - Poland Prof. Mieczysław KACZOROWSKI - Poland Prof. Jouko.J. VUORINEN – Finland Prof. Eduard DORAZIL – Czech Republic Prof. Lubomir BECHNY – Slovakia Prof. Jorge A. SIKORA - Argentina dr Franco ZANARDI - Italy dr Matti JOHANSON – Finland dr Jose R. GUIRIDI - Spain dr Konstantin UZŁOW - Ukraine
ORGANISING COMMITTEE Dr Eng. Jerzy TYBULCZUK - Chairman Dr Eng. Adam KOWALSKI – Secretary Dr Eng. Józef TURZYŃSKI M.Sc.Eng. Krystyna ŁUSZCZKIEWICZ Eng. Marta KONIECZNA M.A. Krystyna BANY-KOWALSKA M.Sc.Eng. Andrzej PYTEL Eng. Janusz CUPIAŁ
PURPOSE AND SUBJECT OF CONFERENCE The purpose of the Conference is to enable the national and foreign research & development centres as well as industrial units to present their latest achievements in developing the heat treatment technology, metallographic examinations, and quality control systems for high-grade ductile iron - ADI in particular.
Therefore you are encouraged to submit your proposals of papers dealing with the following subjects : Foundry New grades of ductile iron, their properties and technologies of production. Methods of spheroidising and inoculation treatments Metals science Electron microscopy. X-ray phase analysis. Quantitative metallography. The techniques of colour etching and their application in metallography. New methods and tools to examine the structure of ductile iron and ADI. Heat treatment Methods of ductile iron heat treatment. Installations. Methods of heat treatment control. Heat treatment parameters. Technologies of ADI fabrication. Quality Quality systems according to ISO and EN Standards used in production of ductile iron and ADI. Control of production process. Statistical quality control.
PRESENTATION OF PAPERS Papers will be presented during the plenary meeting (time for presentation 15-30 minutes). Language of the Conference : English and Polish
INSTRUCTIONS FOR AUTHORS OF THE PAPERS Paper volume : 6-8 pages including tables and figures on A4 paper. Even number of pages. Margins : left - 2.5 cm, right - 2.5 cm, upper - 2.5 cm, lower - 2.5 cm. Write your text in editor Word 6 or Word 97, single line spacing, font Times New Roman 12 pt. The starting line of a new paragraph should be indented by 1 cm.. Figures, photographs and tables should run in the text of the paper. Captions should be typed in bold and centred. Equations should be centred leaving one free spacing above and below the equation. The pages should be numbered consecutively using soft pencil. References : published literature cited in the text should be quoted using a number in square brackets. Abstract : Please start with an abstract of up to 50 words.
SAMPLE OF PAPER FORMAT
On first page (only !) leave free space from the top of 10 single line spacings
TITLE OF PAPER IN BOLD CAPITALS (centred, 14 pt)
space 2 x 1
Name & Surname (bold, centred, 12 pt) Affiliation, e.g. Foundry Research Institute (italics, centred, 12 pt) Place, e.g. Kraków
space 2 x 1 ABSTRACT (bold capitals, centred, 12 pt) space 1 x 1 Text follows - approximately 50 words, indentation of 2.5 cm on the right and left, 12 pt.
space 2 x 1 1. INTRODUCTION (bold capitals, 12 pt) Text justified written in single line spacing. 2. FIRST SUBTITLE (bold capitals, 12 pt) Text justified written in single line spacing. 2.1. Second subtitle (bold, 12 pt) Text justified written in single line spacing. space 3 x 1 REFERENCES (bold capitals, 12 pt) OTHER EVENTS
during the Conference, the universities, industrial plants and companies will have an opportunity to display their products, research methods, manufacturing techniques and computer programmes in the form of short (up to 15 minutes) presentations and/or exhibitions - all those who are willing to take part in the display are kindly requested to agree in advance with the Organising Committee the subject and form of display, the organisers of the Conference also offer the possibility of publishing the ready advertising and information materials in the Conference Proceeding upon previous agreement with the Organising Committee.
Details along with the Conference programme will be circulated early in October 2000 in CIRCULAR NO. 2.
For more information please contact the following persons:
SCHEDULE completed participation forms should be sent by 31 October 2000 technical papers for presentation during the Conference should be sent by 30 September 2000 proposals of promotion during the Conference should be sent by 15 October 2000
REGISTRATION FEE (Note : accomodation is NOT included) full rate: 200 $ reduced rate (for participants presenting papers and posters) 150 $ The cost of promotion is to be agreed.
Please make you cheque payable to : BPH I Oddział w Krakowie Account No. : 10601376-320000033388
Here in Philadelphia we are all looking forward to the upcoming M&M meeting in August. The meeting promises to be the best yet. To make everyone's arrival and traveling as pleasant as possible we recommend you visit the Philadelphia Airport Web site at http://www.phl.org/. Here you will be able to go directly to ground transportation and find the easiest and least expensive ways to get from the airports to the hotels in the city. The ride from the airport to the hotels is approximately 15-20 minutes. Depending on the transportation you choose the prices will range from $8.00 on up. Please feel free to E-mail me with any specific questions you may have and I shall try to assist. We look forward to seeing everyone in August and have a great trip.
I will getting my first SEM/EDS shortly and to obtain the required operating permit from my safety group I need a written procedure for handling LN2. Specifically I need a written procedure for filling the 3 L dewar on the EDS detector from a 50 L dewar mounted on a cart. I plan to make the transfer by using a lab source of N2 to pressurize the 50 L dewar and have obtained all the valves and fittings required to do this from another SEM lab. I would appreciate copies of the procedure. Thanks,
Everett Ramer National Energy Technology Laboratory P.O. Box 10940, Cochrans Mill Road Pittsburgh, PA, USA 15236-0940 Voice: 412-386-4920 FAX: 412-386-4806 ramer-at-netl.doe.gov
The JEOL 840 that I am familiar with (this most likely applies to most JEOL SEM's) sets the tip whenelt distance within a narrow range and as I recall this distance is correct when the auto bias emission current is about 100ua. This makes it possible to fine tune the first cross over with the manual bias control so that at 20kV the emission current would be adjusted with the manual bias control to be about 8-9 and the emission current would be about 10=20uA. The effects of this can be seen viewing the emission pattern. The spot size will be smaller using a high number manual bias number. If the auto bias emission current was say only 50-60uA when the SEM was operated at 1kV accelerating voltage it would be more difficult to fine tune the first cross over spot size because the bias number would be about 0-1 and the emission current would be to low. Increasing the bias number decreases the emission current. If the emission current in auto bias is less than 100uA the whenelt is rotated slightly CW to shorten the distance. Manual bias is the usual way to operate the LaB6 filament while auto bias is usually used with a W filament.
Because the LaB6 source is much brighter than W filament, the first cross over at the source can be fine tuned with the LaB6 cathode which results in smaller spot size on the sample ie. better resolution with sufficient current to have good signal to noise ratio.
If you use a W filament and try to set up manual bias the same way as you would for LaB6 you will see that the signal to noise is worse using high manual bias number (less emission current). The LaB6 filaments are usually operated in manual bias while the W filaments are normally used with auto bias.
On the 840 there are either two accelerating anodes, a high anode for low kV and a low anode for high kV which is the case if you have an elctro static beam blanker installed or a single anode that can be raised for low kV or lowered for high kV operation. The anode in this case snaps into the correct position. Hope this helps.
John Humenansky/Staff Scientist Physical Electronics, Inc. (PHI) 6509 Flying Cloud Drive Eden Prairie, MN 55344 952-828-6387
Hi All, I helped empty on old lab and came across a dozen never-opened bottles of Epon-812 (yes, the real thing). You can well imagine how long they had sat on the shelf. My question to you is....what do I do with it? My gut reaction is to give it to our Life Safety guys to haul away, but if its still good that would be a waste. Short of opening one of the bottles and making a test batch, is there any rule of thumb out there? The bottles have a WPE of 160. They are still in the original plastic wrap and have an EMS label. Maybe Stacie has a thouhgt on this (if she's not overwhelmed with planning for M&M).
Thanks in advance, 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
The Bal-Tec representative is TechnoTrade International. 603-622-5011.
Best,
Angela
At 02:02 PM 06/29/2000 -0400, you wrote:
} I think this query was posted recently but I haven't been able to dredge up } the replies in the list archives - too recent, maybe. } } I'm looking for contact info for a colleague, for Balzers or Bal-tec. He } inherited a B. carbon evaporator model CED 030 (also: 3U-G03-751/132). Can } anyone help out with either contact info for the company, or a copy of the } manual?? We will pay copy/ship charges. } } Thanks, much. } } Ann Hein Lehman } Manager, EM Facility } Trinity College } Hartford CT 06106 } v. 860-297-4289 } f. 860-297-2538 } e. ann.lehman-at-trincoll.edu
--------------------------------------------- Angela V. Klaus
Director, Core Imaging Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
We have used Epon-812 that is over 25 years old with no problem. This is valuable stuff so I hope you do not through it out. Somebody out there will surely take it off your hands. I have enough to last me the rest of my career
At 09:04 AM 06/30/2000 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Gregory W. Erdos, Ph.D. Assistant Director, Biotechnology Program PO Box 110580 University of Florida Gainesville, FL 32611
Hi, Everett. I guess I am glad I am not working for a government anymore. Otherwise I would probably be having to fill out similar forms. It would be nice if the folks that review those forms knew what the issues were. I am always afraid that they will know just enough to be dangerous and may actually require some unhealthy procedures. But perhaps they are sensible and only require that you have something officially in writing to document your best practice.
I found the following post in my files from 4 years ago. I hope Larry does not mind me reposting it. His conclusions are quite interesting.
Warren S.
} Wil's recent comment on the safety hazards of distilled water brought to } mind some peculiar safety regulations here in MD. In reference to liquid N ... } officer listening in will recommend new safety procedures requiring } protective booties! } In the end, we can't legislate common sense, nor can we abdicate } responsibility to those above.
Try getting your safety officer to conduct an experiment:
1. Hold out hand, 2. Pour a small volume of liquid N2 over hand 3. Now the interesting bit - put on a glove, and pour the same quantity of liquid N2 into glove. 4. Phone for ambulance.
The point is that a brief contact causes no problems, but if the contact is continued you get a nasty burn.
Gloves, goggles, masks (and shoes) are actually more dangerous when handling liquid N2 than sandals and no protection. And clothes are actually more dangerous than being naked. Get the safety officer to experiment. With a little persuasion you can probably convince the safety officer that when handling liquid N2, everybody should be naked.
More seriously, bureaucrats, administrators and the inexperienced should talk to somebody who has real knowledge.
-------------------------------------------------------------- Dr. Larry Stoter Technesis 17, Rocks Park Road, Uckfield, E. Sussex, TN22 2AT, United Kingdom Larry-at-teknesis.demon.co.uk --------------------------------------------------------------
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Hi All, I'm asking this on behalf of a colleague, so please bear with me if I don't have all of the details. Does anyone out there have experience with Vectashield as an anti-fade agent for viewing immunofluorescence? Specifically, we are looking at plant tissue in which we are using a secondary antibody conjugated to FITC and have found that the controls (treated only with buffer) show "autofluorescence" after mounting with Vectashield, when they were totally non-fluorescent when mounted in buffer. Granted, the Vectashield did expire last month, but this is something that we've seen before with this product. Should I advise my colleague to just take pictures as quickly as humanly possible because we can't seem to rely on this anti-fade agent? I appreciate any input you may have. Have a great Independence Day weekend (for those of you in the US), Kristen Kristen A. Lennon Cell, Molecular & Developmental Biology Group Department of Botany & Plant Sciences University of California Riverside, CA 92521 kalen-at-citrus.ucr.edu 909-787-4525
} Dear List, } Does any one have the dummies guide to electropolishing?? Some years ago } (3), I had a trouble shooting guide for electropolishing. This list had } about 9 or 10 items, problems /solutions on it and it was very } helpful...but it has found one of those very safe filing places (i.e. I } can't find it!) If any one has access to this list or something similar, } I would appreciate it if you could send me a copy. } } Many thanks in advance. } } Dorrance } PS I haven't been keeping up with the weather reports lately but it's 95 } and clear in beautiful Livermore, California. }
I do not have a "dummies" guide as such, but I do have copies of Bernie Kestel's 66 page report "Polishing Methods for Metallic and Ceramic TEM Specimens". The "polishing" refers to electropolishing. I would be happy to send this out to you if you do not have it already. I also have dozens of other papers dealing with electropolishing and hundreds of papers dealing with specimen preparation. I can email to you a list of the other papers if you'd like to breeze through that. Let me know.
DISCLAIMER: South Bay Technology produces equipment and supplies for specimen prepration and, therefore, has a vested interest in promoting their use.
David Henriks Vice President TEL: 800-728-2233 (toll free in the USA) South Bay Technology, Inc. +1-949-492-2600 1120 Via Callejon FAX: +1-949-492-1499 San Clemente, CA 92673 USA e-mail: henriks-at-southbaytech.com
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy.
Message text written by "McLean, Dorrance" } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} Dear List, } Does any one have the dummies guide to electropolishing?? Some years ago } (3), I had a trouble shooting guide for electropolishing. This list had } about 9 or 10 items, problems /solutions on it and it was very } helpful...but it has found one of those very safe filing places (i.e. I } can't find it!) If any one has access to this list or something similar, } I would appreciate it if you could send me a copy. } } Many thanks in advance. } } Dorrance } PS I haven't been keeping up with the weather reports lately but it's 95 } and clear in beautiful Livermore, California. }
{
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