John: We do around 500 renal biopsies per year and all the sections are mounted on 200 mesh uncoated copper grids. We have an 8 year old Hitachi 7100 and use 60kv. The majority of the glomerulus can be viewed with the 3-4 serial sections lying randomly across the grid bars. We do not need a picture of the whole glomerulus, rather most pictures are between 3,000 and 10,000X. Dr. Tibor Nadasdy is the renal pathologist and decided last year that all our renal biopsies would be captured with the digital camera onto a computer and sent up to him via a network to his computer. So, at the present time we use very little EM film. He diagnoses each biopsy and e-mails representative digitized images to the nephrologists.
Karen L. Jensen, M.S. Project Manager & Associate Scientist Electron Microscopy Research Core
-----Original Message----- } From: "JHoffpa464-at-aol.com"-at-sparc5.microscopy.com [mailto:"JHoffpa464-at-aol.com"-at-sparc5.microscopy.com] Sent: Friday, March 30, 2001 2:20 PM To: microscopy-at-sparc5.microscopy.com
Hello -
Has anyone out there tried to adhere xylem sections to pre-coated microscope slides like Fisher Probe-On Plus slides? I've tried and had a zero percent section retention. I can imagine that this is because of the scarcity of live cells (and plasma membranes). I've tried slow air drying and various temperatures on the slide warmer. The sections are 15 microns, and are from fixed (buffered paraformaldehyde) but not embedded samples. I'm about to try Poly-L-lysine and amino-acyl silane treated slides, but I'm not too hopeful because they (at least Poly-L) rely on the same positively-charged surface principle. I want to avoid gelatin or albumin subbing because I'm treating sections with protease, and also want to minimize background staining. Any tips would be greatly appreciated.
Rachel
****************************************** Rachel Spicer Biological Laboratories 3119 Organismic and Evolutionary Biology Harvard University 16 Divinity Avenue Cambridge, MA 02138
Rishi Raj wrote: ============================================================ We have just acquired a used microscope made by ISI Inc. (their model alpha - 1980). Would anyone please have information on how we can obtain spare parts, filaments etc., for this machine. Many thanks for your help... ============================================================ The business of the ISI SEM's is now being handled by
Aspex Instruments LLC Formerly: RJ Lee Instruments Ltd. 175 Sheffield Drive Delmont, PA 15626 USA Tel: 1-724-468-5400 Fax: 1-724-468-0225 E-mail: pssales-at-rjleeinst.com
The former manager of the SEM operation when it was still ISI, Michael McCarthy, is now with Aspex. Michael might be the single most knowledgeable person in terms of spare parts for the column and vacuum system. Several of the main suppliers of consumables, like SPI Supplies, also offer filaments, new and retipped, apertures, and the other items of that nature you would be needing to maintain the microscope.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
I desperately want to be a materials researcher/Electron Microscopist. We have a Newly established Electron Microscope Laboratory. We need knowledge and skills. Please assist for a scholarship/suport as we have no funds, nor courses of the like at our University.
Hello everybody, We have an JEOL JXA 8600 EPMA in our Institute, with three WDS spectrometers. We are planning to buy an other one, and we where thinking about the H-type x-ray spectrometer.
Is there anybody in the list who has experience with this kind of spectrometers that could give me some information about them (their performance in general and also comparatively to standard spectrometers, etc.)?
Thanks
Laura Hernandez Laura Hernandez Laboratorio Microsonda Electronica Instituto GEA Universidad de Concepcion Casilla 160C Concepcion CHILE
Try using Vectabond treated slides. vectabond is available from vector laboratories. We had quite good results with leaf, stem, root sections sticking to these slides. Vector lab: 1-800-227-6666
Good luck,
Soumitra
I have no financial interest in Vector Laboratories.
On Sat, 31 Mar 2001, Rachel Spicer 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 - } } Has anyone out there tried to adhere xylem sections to pre-coated } microscope slides like Fisher Probe-On Plus slides? I've tried and had a } zero percent section retention. I can imagine that this is because of the } scarcity of live cells (and plasma membranes). I've tried slow air drying } and various temperatures on the slide warmer. The sections are 15 microns, } and are from fixed (buffered paraformaldehyde) but not embedded samples. } I'm about to try Poly-L-lysine and amino-acyl silane treated slides, but } I'm not too hopeful because they (at least Poly-L) rely on the same } positively-charged surface principle. I want to avoid gelatin or albumin } subbing because I'm treating sections with protease, and also want to } minimize background staining. Any tips would be greatly appreciated. } } Rachel } } } } } ****************************************** } Rachel Spicer } Biological Laboratories 3119 } Organismic and Evolutionary Biology } Harvard University } 16 Divinity Avenue } Cambridge, MA 02138 } } (617) 496-3580 (phone) } (617) 496-5854 (fax) } spicer-at-oeb.harvard.edu } ****************************************** } } }
You can do a brightness leveling image process with a number of a standard packages.
The procedure is simply to do a Gaussian blur that gives you the slowly varying component of the background and subtract that from your original image.
-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: Feng Wu [mailto:fwu-at-bgumail.bgu.ac.il] } Sent: Wednesday, March 28, 2001 6:14 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: adjust the intensity of the image } Sensitivity: Confidential } } } -------------------------------------------------------------- } ---------- } 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/MicroscopyLists } erver/FAQ.html } } } } -------------------------------------------------------------- } ---------. } } } Hi, All, } I take some diffraction contrast pctures. As the thickness of } the samples } changes sharply the brightness of the images is not uniform. } Does someone know } any software to adjust the brightness for the whole image? } Thanks inadvance. } Best regards. } Feng } ********************************************** } Dr. Feng Wu } Dept. of Materials Engineering } Ben-Gurion University of the Negev } Beer-Sheva 84105, Israel } } voice 972-8-6461473 } } fwu-at-bgumail.bgu.ac.il } ********************************************** } } }
I'm posting this message for a colleague; as always, I appreciate your help. Alice.
Alice Dohnalkova Environmental Microbiology Battelle, PNNL Richland, WA (509) 372-0692
} My graduate student has made TEM images of our plasma polymerized aniline } films. The films seem to have a "cauliflower" structure that could } probably be described as a fractal pattern. Have you ever made plasma } polymerized aniline films and if so did you see a cauliflower structure? } Your comments would be helpful. Thanks. } Pat } } Patrick D. Pedrow, pedrow-at-eecs.wsu.edu, www.eecs.wsu.edu/~pedrow
} Greetings, } I have found that polyethlyene-imine (PEI) is much stickier } than poly-lysine. I have no idea about the resin you mentioned, but } PEI is sticky stuff. It comes as a liquid. I make a 0.1% soloution in } ddwater, which I freeze in aliquots. Then I keep a working one in the } fridge. I coat coverslips in the stuff by floating them on a drop of } the PEI solution for about 10 sec, and then blotting off the excess } and letting them air dry. In that conditions, the coated 'slips are } good for at least months. } } Hope this helps, } Tobias Baskin }
Tobias:
Would this be Sigma catalogue number P-3143?
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 **********************************************
I am looking for a X-ray radiation detector used on TEMs. Currently we have a Geiger counter that is only sensitive to low energy X-ray. The purpose of the new detector is to sense high energy X-ray leakage from a 200 kV scope.
Any suggestion or clue will be greatly appreciated. Please contact off-line.
Dear Haifeng, The x-ray monitors on our HVEM are wrapped in a metalic sheath so they will be sensitive to the brehmsstrahlung spectrum of 1.2 MeV electrons. I don't think that they could be easily connected to your scope, but you might look into making a similar sheath for your existing Geiger counter. Good luck. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
I have worked with CAMECA SX50 with the horizontal spectrometer. It was helpful when working with fractures, for example for identification of nonmetallic inclusions. Another it's advantage was that it was the only spectrometer with all 4 crystals, so it had more flexibility in maps or line scans acquisition. In quantitative analysis I did not use it both major and trace elements acquisition and never got any problems with its performance.
Vladimir Dusevich
-----Original Message----- } From: Laura Hernandez To: Microscopy-at-sparc5.microscopy.com Sent: 4/2/01 7:41 AM
Hello everybody, We have an JEOL JXA 8600 EPMA in our Institute, with three WDS spectrometers. We are planning to buy an other one, and we where thinking about the H-type x-ray spectrometer.
Is there anybody in the list who has experience with this kind of spectrometers that could give me some information about them (their performance in general and also comparatively to standard spectrometers, etc.)?
Thanks
Laura Hernandez Laura Hernandez Laboratorio Microsonda Electronica Instituto GEA Universidad de Concepcion Casilla 160C Concepcion CHILE
I am trying to determine a method to create or embed points of reference on an SEM sample. To give some background, the samples are sections of microprocessor packages that are placed in a modified stage with a three-point bend fixture. What I'm trying to do is monitor the movement of points on the specimen surface as the load is increased. I tried using the spot mode on the scope to see if I could remove some of the sputter coat, since most of the material underneath is non-conductive, but that was unsuccessful.
Imaging will most likely be done between 300 and 1000x. I need to have multiple reference points on the screen at one time, since I will be measuring relative displacements. The reference points do not need to be distributed in a uniform pattern.
I have tried applying some powder to the surface, since I read about this being done before, but the results were not acceptable. I may just be using the wrong type of powder, but I haven't been able to find out what kind of powders would work best
So, any ideas on how I may accomplish this?
Thanks,
Norman Kay Graduate Student AME Dept. The University of Arizona
I want to put a (cheap) video camera onto the optical microscope of my JEOL 840.
I want to leave the eyepiece lens on, so that users can have the option of easily removing the camera to use the microscope conventionally.
I have tried presenting several different models of CCD video cameras up to the eyepiece, both with and without the camera lens attached, but none gives me anything better than a smallish bright circle in the centre of the (black) field of view.
I presume that I need some sort of intermediate lens, but my understanding of physical optics has largely evaporated over the years.
Can someone point me towards a suitable text or other information source?
thanks
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
} From: "Ritchie Sims" {r.sims-at-auckland.ac.nz} } I want to put a (cheap) video camera onto the optical microscope of } my JEOL 840. } } I want to leave the eyepiece lens on, so that users can have the } option of easily removing the camera to use the microscope } conventionally. } } I have tried presenting several different models of CCD video cameras } up to the eyepiece, both with and without the camera lens attached, } but none gives me anything better than a smallish bright circle in } the centre of the (black) field of view. } } I presume that I need some sort of intermediate lens, but my } understanding of physical optics has largely evaporated over the } years. } } Can someone point me towards a suitable text or other information } source? } } thanks } } rtch
A CCD camera without a lens looking into and eyepiece usually has the opposite problem of having too much magnification. The image coverage of the image on the CCD camera can be increased when no lens is present on the camera simply by moving the camera further away from the eyepiece. My set up uses a 2.6x eyepiece for the video camera and a 10 x eyepiece to view the distance between the 2.6 eyepiece and the CCD element is about 2 inches or a little more and I have almost twice the magnification on the CCD camera as I see through the 10X eyepiece.
So just adding a spacer between your camera and your eyepiece should solve your problem.
Gordon Gordon Couger gcouger-at-couger.com Stillwater, OK www.couger.com/gcouger
If you have coat your specimen with a thin carbon layer for conduction then add another layer of gold through a TEM grid as a mask you should be able to see the grid bars. Check out your local EM supplier's catalogue for the most suitable grid design.
Good luck, Ron
On Mon, 2 Apr 2001 15:48:48 -0700 SEM Machine {SEM-at-ACATC.AME.Arizona.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 am trying to determine a method to create or embed points of reference on } an SEM sample. To give some background, the samples are sections of } microprocessor packages that are placed in a modified stage with a } three-point bend fixture. What I'm trying to do is monitor the movement of } points on the specimen surface as the load is increased. I tried using the } spot mode on the scope to see if I could remove some of the sputter coat, } since most of the material underneath is non-conductive, but that was } unsuccessful. } } Imaging will most likely be done between 300 and 1000x. I need to have } multiple reference points on the screen at one time, since I will be } measuring relative displacements. The reference points do not need to be } distributed in a uniform pattern. } } I have tried applying some powder to the surface, since I read about this } being done before, but the results were not acceptable. I may just be using } the wrong type of powder, but I haven't been able to find out what kind of } powders would work best } } So, any ideas on how I may accomplish this? } } } Thanks, } } Norman Kay } Graduate Student } AME Dept. } The University of Arizona } } }
---------------------- 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
The magnifications you are using present the problem. I'm sure that you are also wanting as much resolution as possible from image processing of the resultant images.
Producing a non-conductive spot on the sample is a good idea, as it should stand out well, particularly during a slow record mode scan.
My vote - copier or laser printer toner. Very small particle size, non-conductive plastic composition. Also, once the powder is sprinkled on, it can be adhered to the surface with a little heat to ensure that it doesn't move around.
On Monday, April 02, 2001 5:49 PM, SEM Machine [SMTP:SEM-at-ACATC.AME.Arizona.edu] wrote: } } } I am trying to determine a method to create or embed points of reference on } an SEM sample. To give some background, the samples are sections of } microprocessor packages that are placed in a modified stage with a } three-point bend fixture. What I'm trying to do is monitor the movement of } points on the specimen surface as the load is increased. I tried using the } spot mode on the scope to see if I could remove some of the sputter coat, } since most of the material underneath is non-conductive, but that was } unsuccessful. } } Imaging will most likely be done between 300 and 1000x. I need to have } multiple reference points on the screen at one time, since I will be } measuring relative displacements. The reference points do not need to be } distributed in a uniform pattern. } } I have tried applying some powder to the surface, since I read about this } being done before, but the results were not acceptable. I may just be using } the wrong type of powder, but I haven't been able to find out what kind of } powders would work best } } So, any ideas on how I may accomplish this? } } } Thanks, } } Norman Kay } Graduate Student } AME Dept. } The University of Arizona } } } }
Allen R. Sampson, Owner Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174 voice 630.513.7093 fax 630.513.7092
My collegue C. Ulhaq want to know who practice Lorentz Microscopy on TEM (in Europe particulary).
The questions would be : which kind of microscope you use, do you use special polar pieces, and did you buy it or were they "home made". Same question about the sample holder. What are the max magnification accessible ? We have a ABT Topcon 002B, with the the possibility to change the polar pieces.
You can answer direct to my collegue (corinne.ulhaq-at-ipcms.u-strasbg.fr), or on the list.
J. Faerber IPCMS-GSI (Institut de Physique et Chimie des Matériaux de Strasbourg Groupe Surface et Interfaces) 23, rue de Loess 67037 Strasbourg CEDEX France
Thanks for the replies and the advice that you have offered.
I have Some additional information. I am a BSc.[Physics, Mathematics] holder. Currently I am working at the E.M laboratory of the university of Dar es Salaam as a supporting staff, and I am looking for the opportunity to pursue an MSc. and hence a Ph.D that will qualify me to research fellow. Funds are a hindrance. We have one old ZEISS9S2 TEM, and a new LEO 910 analytical TEM. An MSc. course in TEM or a research in metals/ceramics will do.
Hello, We have a client who needs to embed a cell culture in paraffin. In our lab we embed cultures in agarose and then embed for TEM. We do not have experience embedding paraffin. My question is, since xylene is used for paraffin how do you keep the culture from dispersing. Any suggestions? Thank you
Karen Kelley Senior Electron Microscopist University of Florida ICBR Electron Microscopy Core Lab Box 118525 Gainesville Florida Lab: 352-392-1184 fax: 352-846-0251 email: klk-at-biotech.ufl.edu http://www.biotech.ufl.edu/~emcl/staff/karenpage.html
I have not done Lorentz microscopy (here LEM) in a long time and I am not very familiar with the configuration of the Topcon 002B. This is what I remember ,for LEM to work the sample has to be outside the strong magnetic field of the OL pole piece. In the good old days, we used a modified top entry-type specimen holder which was longer than usual so that the specimen would sit below its normal position. In addition, the IL electronics had to be modified so that focus could still be attained when the specimen was in this lower position. We normally used magnifications of up to about 20 KX if I recall.
The second way of doing this is to essentially turn off the objective lens , but you have to be able to focus with the IL . Some of the newer scopes might not be able to do this without modifications to the electronics. In our case all the modifications were done by the manufacturer.
Hope this helps
Jordi Marti
-----Original Message----- } From: Faerber Jacques [mailto:Jacques.Faerber-at-ipcms.u-strasbg.fr] Sent: Tuesday, April 03, 2001 3:25 AM To: Microscopy Society of America
Hi everyone, We are working with suspension cells (Jurkats or H9s) and we need a protocol to prepare them for fluorescent microscope (both fixed and live cell imaging). Thanks in advance for your help.
Asli Oztan
asost2-at-pitt.edu University of Pittsburgh Molecular Virology and Microbiology
You can try spinning (centrifuging) the cells down so they become a packed ball of cells, then carefully put them into a bag (i think they are nylon bags) for processing and subsequently into paraffin block.
Frank Lee
Karen Kelley 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, } We have a client who needs to embed a cell culture in paraffin. In our lab } we embed cultures in agarose and then embed for TEM. We do not have } experience embedding paraffin. My question is, since xylene is used for } paraffin how do you keep the culture from dispersing. Any suggestions? } Thank you } } Karen Kelley } Senior Electron Microscopist } University of Florida } ICBR Electron Microscopy Core Lab } Box 118525 Gainesville Florida } Lab: 352-392-1184 fax: 352-846-0251 } email: klk-at-biotech.ufl.edu } http://www.biotech.ufl.edu/~emcl/staff/karenpage.html
Dear Norman, When we had a similar problem of creating a reference for studying crack growth, we used the gold-evaporated grid method, as Ron Doole mentioned. However, this gave problems because the grid is uniform, so after you've traveled a little way, you had no unique reference to keep you placed. We solved this by also puting a drop of latex sphere suspension on the surface, before sputter coating. This is a suspension of latex spheres of specific size, I believe we used one micron, but you can use a size suitable to your magnification. The suspension dries to form a random pattern of dots that can be compared in photos. You amy have to experiment with the concentration of spheres to get the right coverage. We were lucky enough to have a researcher making latex spheres who gave us some of her duds, but these suspensions can be purchased. I hope this helps. At 03:48 PM 4/2/01 -0700, you wrote: } } I am trying to determine a method to create or embed points of reference on } an SEM sample. To give some background, the samples are sections of } microprocessor packages that are placed in a modified stage with a } three-point bend fixture. What I'm trying to do is monitor the movement of } points on the specimen surface as the load is increased. I tried using the } spot mode on the scope to see if I could remove some of the sputter coat, } since most of the material underneath is non-conductive, but that was } unsuccessful. } } Imaging will most likely be done between 300 and 1000x. I need to have } multiple reference points on the screen at one time, since I will be } measuring relative displacements. The reference points do not need to be } distributed in a uniform pattern. } } I have tried applying some powder to the surface, since I read about this } being done before, but the results were not acceptable. I may just be using } the wrong type of powder, but I haven't been able to find out what kind of } powders would work best } } So, any ideas on how I may accomplish this? } } } Thanks, } } Norman Kay } Graduate Student } AME Dept. } The University of Arizona } 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
Can someone identify a supplier for small amounts of methlyamine tungstate powder (used for negative staining in TEM)? I have tried EMS, Ted Pella, and Polysciences (the supplier of our original decade-old vial). Robert J. Palmer Jr., Ph.D. Natl Inst Dental Craniofacial Res - Natl Insts Health Oral Infection and Immunity Branch Bldg 30, Room 308 30 Convent Drive Bethesda MD 20892 ph 301-594-0025 fax 301-402-0396
I'm trying to get a handle on the value of an old Auger/ESCA spectrometer. The model is PHI 558, by Perkin-Elmer. It was built around 1985 or 86, and the electronics are quite old, but it's intact and fully functional. It also has a LEED detector and has been upgraded over time. A partial list of parts is included below. Does anyone have an idea what this might be worth?
Thank you, Ellen Carrillo-Heian
emheian-at-engr.ucdavis.edu Dept. of Chem. Eng. and Mat. Sci. UC Davis Davis, CA 95616 USA ---------------------- Partial list of components:
} 32-010 Lock In Amp } 20-0275 Electron Multiplier Supply } 32-095 X Ray Source Control } 22-040 DC Power Supply } 16-020 Heat Exchange / Deionizer } 20-805 Analyzer Control } 32-100 Electron Multiplier Supply } 11-065 Ion Gun Control } 20-115 Ion Gun Control } 11-010 Electron Gun Control } 11-055 ESCA / Auger System Control } 11-500A Auger System Control } Inficon 012-214 } 04-303 Differential Ion Gun } 04-548 Dual Anode X-ray Source } 15-255 GAR Precision Electron Energy Analyzer } Ultek DI Pump } 218075B-26 UHV Instruments (Tag in front of the screw/bellows assembly, on } frame.) } } And a 386 Compaq controlling it. } } The chamber has the dual chamber translation set up. (With the 4 ft } screw/bellows) }
We do this using contamination spots from the microscope itself. We put down a grid array of spots using our Isis system to control the stage and spot positons. Focus the beam to a spot and let is sit there for a minute or so and a contamination spot should appear. We are using electropolished aluminum. I dont know if it will work for your material but its worth a try Then we strain our material and look at it again. Works well.
I would like to do high resolution TEM on spherical, semiconductor nanoparticles. The diameters of these particles range from 1 to 10 nm. Our goal is to acquire good digital images of these particles so that we can size them in house. Please let me know if you can help and how much you charge per sample or per hour. Your help will be greatly appreciated.
Dear Dr. Palmer, Nanoprobes sells the methylamine tungstate you desired under the name "NanoW". It is an excellent negative stain. They also make methylamine vanadate, a similar, but lower atomic number stain they call "NanoVan". More information is at www.nanoprobes.com. J. Hainfeld
Dr. James F. Hainfeld Brookhaven National Laboratory Biology Dept. Bldg. 463 Upton, NY 11973 USA Tel. 631-344-3372 Fax. 631-344-3407 email: hainfeld-at-bnl.gov website: http://bnlstb.bio.bnl.gov/biodocs/stem/stem.html
Generally, equipment depreciates at the rate of 30% per year on the balance. In other words, an SEM with an initial cost of 100K is worth 70K after the first year, 49K the second year, 34.3k the third year, etc.
These numbers are based upon the experience I have had with used equipment sales during the past five years. Other allowances are made for equipment that has been abused, etc. Strangely enough, accessories add little to the resale value of equipment.
I am sure there are others who would differ as it would not fit into their accounting procedures but my experience has been that scientific equipment depreciates more than computers.
Regards,
Earl Weltmer
I have no financial interest in this thread only experience.
----- Original Message ----- } From: "Ellen Carrillo-Heian" {emheian-at-engr.ucdavis.edu} To: "microscopy" {Microscopy-at-sparc5.microscopy.com} Sent: Tuesday, April 03, 2001 11:25 AM
Hi everybody (again),
I am looking for a program that allows me to quantify a given group of nanoparticles that have various shapes and sizes. These particles range from 1 to 10 nm in diameter. Your help will be greatly appreciated. Thank you.
We sell this as a 2 % aqueous solution (suitable for use directly) - the product is called "Nano-W." It's listed on our web site catalog under "Negative staining."
Regards,
Rick Powell
***************************************************************************************** Richard D. Powell * rpowell-at-nanoprobes.com * www.nanoprobes.com NANOPROBES, Incorporated 95 Horse Block Road, Yaphank, NY 11980-9710, USA
Sign up for our newsletter: http://www.nanoprobes.com/Newsletter.html *****************************************************************************************
What kind of computer depreciates (looses value) slower than scientific equipment? A regular PC or Mac drops by over 50% the first year. After an additional six months, the system worth next to nothing. But of course, the new and improved model is $2K or more. Either way, the standard IRS depreciation schedule for computers and scientific equipment is five years. I would say that this is more appropriate for scientific equipment than it is for computers. But YMMV.
gary g.
At 05:16 PM 4/3/2001, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
This is an interesting thread. Our unit is supposed to operate at "full cost recovery", and as such we are charged depreciation of the instruments (funded from a central equipment grant), which are depreciated over 15 years. Only computers are depreciated over 3 years. If anyone else is aware of some vaguely standard depreciation times for TEMs, SEMs, confocal, light microscopes, I'd appreciate hearing about this.
Thanks, rosemary
Rosemary White Microscopy Centre CSIRO Plant Industry GPO Box 1600 Canberra, ACT 2601 Australia
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Rosemary said - } This is an interesting thread. Our unit is supposed to operate at "full } cost recovery", and as such we are charged depreciation of the instruments } (funded from a central equipment grant), which are depreciated over 15 } years.
FWIW, I understand that that our organization (A Canadian federal government one) also depreciates this kind of major capital equipment over 15 years. (Except, apparently, helicopters - our Navy is still using ones which are, literally, older than most of the pilots flying them....)
Frank Thomas Geological Survey of Canada (Atlantic) Bedford Institute of Oceanography Dartmouth, Nova Scotia
John Russ recently demonstrated an adaptive equalization technique for us that works quite well. Basically it is histogram equalization over a local area of the image that is then stepped over the whole image. He has implemented this method in his IP Toolkit and Fovea Pro packages that plugin to PhotoShop. It is also described in The Image Processing Handbook, 2nd edition on page 222.
Just a satisfied customer...
Henk Colijn
At 11:42 AM 4/2/01 -0400, Walck, Scott D. wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hendrik O. Colijn colijn.1-at-osu.edu Campus Electron Optics Facility Ohio State University (614) 292-0674 http://web.ceof.ohio-state.edu Fools are pleased when they discover error. The wise are pleased when they discover truth.
An interesting thread indeed! We are in the midst of surplusing out a lot of used histology equipment, and find that there are two different methods of determining value. Depreciation of equipment for tax purposes is done according to state/federal tax laws--and I think that means after 5 or 10 years (the time seems to change, and I can never keep up with) the _tax_ value is zero. However. When it comes to disposing of the equipment, our business folks are using a different "book value" that has values significantly greater than current market value. So.... Draw your own conclusions.
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc.
On Wed, 4 Apr 2001 15:47:39 +1000, Rosemary White 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 | -----------------------------------------------------------------------. | | | This is an interesting thread. Our unit is supposed to operate at "full | cost recovery", and as such we are charged depreciation of the instruments | (funded from a central equipment grant), which are depreciated over 15 | years. Only computers are depreciated over 3 years. If anyone else is | aware of some vaguely standard depreciation times for TEMs, SEMs, confocal, | light microscopes, I'd appreciate hearing about this. | | Thanks, | rosemary | | | Rosemary White | Microscopy Centre | CSIRO Plant Industry | GPO Box 1600 | Canberra, ACT 2601 | Australia | | phone 61-2-6246 5475 | fax 61-2-6246 5000 | email r.white-at-pi.csiro.au | | |
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc. 900 Rigdebury Road Ridgefield, CT 06877 203-798-5448
_______________________________________________________ Send a cool gift with your E-Card http://www.bluemountain.com/giftcenter/
I think that CSIRO is on the high side. In the Canadian government 10 years seems to have been the semi-official standard for as long as I can remember. In my lab, though, we informally think of two 'effective lifetimes', 7 years or thereabouts for the TEM and SEM, and 10 for the other beam instruments (EPMA, SIMS, XPS and SAM). Furthermore, we regard these as upper limits for two reasons:
- vendors these days are operating on ever-tighter parts inventories, thus a beam instrument may have some good years left in it, but you find you can't get a crucial part any longer. This happened to us recently with our 12 year old Cameca SIMS, when a flight tube developed ultrafine cracks and we discovered that they are not kept in stock any more. After a lot of arm-twisting and calling of favors, we tracked down one of the few remaining ones in Europe, otherwise we would have been down for 3 months awaiting a custom-built one.
- we do a lot of work with the private sector, some on a contract basis. They often come to us to get state-of-the-art data quality which has been collected in a timely fashion. The first is what usually gets their attention in the first place, while the second is crucial to keeping their attention.
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: Rosemary White } Sent: Wednesday, April 04, 2001 1:47 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: equipment depreciation } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } This is an interesting thread. Our unit is supposed to operate at "full } cost recovery", and as such we are charged depreciation of the instruments } (funded from a central equipment grant), which are depreciated over 15 } years. Only computers are depreciated over 3 years. If anyone else is } aware of some vaguely standard depreciation times for TEMs, SEMs, } confocal, } light microscopes, I'd appreciate hearing about this. } } Thanks, } rosemary } } } Rosemary White } Microscopy Centre } CSIRO Plant Industry } GPO Box 1600 } Canberra, ACT 2601 } Australia } } phone 61-2-6246 5475 } fax 61-2-6246 5000 } email r.white-at-pi.csiro.au } } }
I am processing samples of white fat for TEM and have been having trouble with a dense precipitate that covers the cytoplasm. Nuclei and blood vessels are relatively unaffected. So far I've tried 3 different fixatives: Trump's and 2.5% glut/2% para in either 0.1M cacodylate of 0.1M phosphate buffer. The samples have been osmicated for one hour and embedded in either Spurr's resin or Epon. The precipitate is present in unstained sections as well as those stained with UA alone or UA and lead. Treatment with .5N HCl for 2 minutes alleviates the problem somewhat but bleaches the sections so much that they are very difficult to see. If anyone has encountered similar problems and has suggestions for me I would be most grateful.
Thanks in advance
Germaine G. Boucher TEM Lab Pfizer Global Research and Development Groton, CT
We are looking to replace a very old EDS system on our Hitachi S-2460N variable pressure SEM. I would like to know about your preferences for and experiences with different companies, both good and bad. Please reply directly to me. If anyone else is interested, I could submit a summary to the list after I've collected all the responses.
Thanks in advance.
Jean Ross Central Microscopy Research Facility University of Iowa
I have two questions concerning service contracts on confocals. Let me start by saying i have had a confocal for about 8 years and would never consider going without one. I have a Biorad 2000 going off warranty and need to make a decision.
First question: Biorad no longer guarantees a response time - they now promise to get to you as fast as they can but no longer promise a 48 or 72 hr response. Have other confocal manufacturers done this also?
Second question: My university is pushing replacing service contracts with "insurance" contracts with a major vendor who then pays for a service visit from the manufacturer on an hourly basis. All parts, travel, service repair time, etc are covered at a price that is typically 75% less than the manufacturer's service contract. They guarantee the price and coverage for 3 years. Personally I don't know how they could make money on this deal since we average a fair number of visits and spare parts (e.g. lasers) in a typical year. Does anyone have experience with this type of situation with confocals? The company the University is dealing with is CIC but there are several other ones out there.
Thanks for any input. -- 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)
dear listservers.... i need some advice.... we use Permount as our mountant...however, as we cut our slides extremely thick (120 micra) for our staining method (Golgi-impregnations of neurons), it often takes 2-3 weeks for the Permount to dry sufficiently so that we can actually use the slides without it getting on our microscope stage, or the coverslip moving around under our oil-immersion lenses.... so....my question is --- does any microscope maven out there know if there is anything that can be done to accelerate the drying/hardening the Permount....??? many thanks for any help.... regards, Ron Mervis ~~~~~~~~~~ Ronald F. Mervis, Ph.D. Neuro-Cognitive Research Laboratories 2109 West Fifth Avenue Columbus, Ohio 43212 USA ~~~~~~~~~~~~~~~~~~~~~~~~~~~~ ..independent nonprofit contract laboratories dedicated to quantitiative neurostructural analysis to promote our knowledge and understanding of human neurological diseases, neurodegeneration, and neuroplasticity.... ~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Tel: (614)-486-6404; lab: (614)-486-6080 Fax: (614)-486-6020 e-mail: RonMervis-at-aol.com (or) RonMervis-at-Neuro-Cognitive.org ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ "...can the human soul be glimpsed through a microscope? Maybe, but you'd definitely need one of those very good ones with two eyepieces." - Woody Allen
{HTML} {FONT FACE=arial,helvetica} {FONT SIZE=2} dear listservers.... {BR} i need some advice.... {BR} we use Permount as our mountant...however, as we cut our slides extremely {BR} thick (120 micra) for our staining method (Golgi-impregnations of neurons), {BR} it often takes 2-3 weeks for the Permount to dry sufficiently so that we can {BR} actually use the slides without it getting on our microscope stage, or the {BR} coverslip moving around under our oil-immersion lenses.... {BR} so....my question is --- does any microscope maven out there know if there is {BR} anything that can be done to accelerate the drying/hardening the {BR} Permount....??? {BR} many thanks for any help.... {BR} regards, {BR} Ron Mervis {BR} ~~~~~~~~~~ {BR} Ronald F. Mervis, Ph.D. {BR} Neuro-Cognitive Research Laboratories {BR} 2109 West Fifth Avenue {BR} Columbus, Ohio 43212 USA {BR} ~~~~~~~~~~~~~~~~~~~~~~~~~~~~ {BR} ...independent nonprofit contract laboratories dedicated to quantitiative {BR} neurostructural analysis to promote our knowledge and understanding of human {BR} neurological diseases, neurodegeneration, and neuroplasticity.... {BR} ~~~~~~~~~~~~~~~~~~~~~~~~~~~~ {BR} Tel: (614)-486-6404; lab: (614)-486-6080 {BR} Fax: (614)-486-6020 {BR} e-mail: RonMervis-at-aol.com (or) RonMervis-at-Neuro-Cognitive.org {BR} ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ {BR} "...can the human soul be glimpsed through a microscope? Maybe, but you'd {BR} definitely need one of those very good ones with two eyepieces." {BR} - Woody Allen {BR} {/FONT} {/HTML}
I have seen some beautiful SEM pictures of semiconductor interconnect lines (Copper and aluminum) in which the dielectric material (Silicon dioxide I assume) has been completely removed. Can someone tell me what was used to remove the dielectric without affecting the metallization?
Hello friends, I have a gold bronze composition material coming in for imaging by SEM, x-ray mapping, and optical microscopy. I could us a sample prep recommendation, particularly for a etchant or so I think. This is primarily a failure analysis project in which we want to clearly observe and differentiate grain boundaries. Recommendations would be greatly appreciated.
thanks, Bruce Brinson Optical Analyst Rice University
Germaine, If you don't rinse in buffer well enough after the initial fixation , the glut will form a precipitate with osmium. Try washing 4x or 5x for 15 minutes each between glutaraldehyde and osmium. Good luck,
At 10:34 AM 4/4/01 -0400, Boucher, Germaine G wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Mary Gail Engle Sr. Research Laboratory Manager Electron Microscopy & Imaging Facility Health Sciences Research Bldg. 001 University of Kentucky Lexington, KY 40536-0305
Thanks for all the recommendations for suppliers of cameras and interfaces, but what I was wanting was a pointer to a text or somesuch from which I could figure out myself what I need.
There must be someone in this learned and experienced community who's been there and done that, isn't there?
"That", for those who may have missed my first post, being the problem of how to work out what sort of intermediate lens would be needed to interface a small cheap CCD video camera (or a webcam) so that it gives a good image when looking into the existing eyepiece lens of a given optical microscope (in this case the OM of a JEOL 840 SEM).
thanks
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
Dear colleagues, I would like to buy a digital camera for my Axiolab Zeiss microscope. Unfortunately I am a bit confused in the amount of available data. I would need a digital camera of the resolution that matches the quality of film cameras in order I need not scan photos or negatives. Could you please be so kind and give me a piece of advice? I have Olympus Camedia 3030 with 3,34 mil pixels. Will this camera and resolution do or do I need to buy another one? If yes of what resolution and type? I will be very indebted for an advice because I receive often contradicting information for different distributors and I am not much wise about it. Many thanks in advance. With best wishes Jiri Kalvoda Department of Geology and Paleontology Masaryk University Kotlarska 2 61137 Brno Czech republic
It is easily done with a plasma etch using CF4 + O2.
The top layer of "glass" is typically silicon nitride over silicon dioxide or in earlier devices, it is boron phosphor silicon glass (BPSG).
I have some colorized shots on my web site at:
http://photoweb.net
More get added and some get swapped over time.
gary g.
At 11:48 AM 4/4/2001, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Rod McCabe wrote: ======================================================= I have seen some beautiful SEM pictures of semiconductor interconnect lines (Copper and aluminum) in which the dielectric material (Silicon dioxide I assume) has been completely removed. Can someone tell me what was used to remove the dielectric without affecting the metallization? ======================================================= While HF and Q-Tips can be used to swab off the SiO2, the better way (in our opinion) is with reactive plasma etching, using CF4 as the reactive gas. This way the layer is removed in a way that does not disrurb, for example, corrosion product that might have formed underneath. If you use the wet chemical approach, you can dissolve and swab away features of interest, such as corrosion product. And you have removed that which you might otherwise have been able to analyze with EDS or even Auger.
Several manufacturers offer table top plasma etchers, SPI Supplies being one of them. You can find information about the SPI Plasma Prep™ II on our 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.2spi.com ############################ ==================================================
I am writing for comments -- off-list please -- about the CleanZone LF clean air workstation manufactured by IQAir, which, I am told, is used in Switzerland and Germany, but only recently has been introduced in the United States. With thanks,
James Martin Orion Analytical, LLC www.orionanalytical.com martin-at-orionanalytical.com
A colleague has asked for recommendations for setting up a digital darkroom (fun to spend someone else's money!). This person would benefit from a really good scanner that could deal with prints, large format negatives (4"x5", 3.25"x4") as well as 35 mm slides. At one time I looked into an Agfa Duoscan T2500. Do any of you have an opinion about this or other suitable scanners?
I know this subject comes up regularly, but I don't feel bad about introducing it again, since technology evolves so quickly!
Mahalo, Tina
http://www.pbrc.hawaii.edu/bemf/microangela **************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
thanks to everyone for the advices about all type of glues for tripod polishing. I tried to make some sort of summary from the different answers I have received. I would be happy if it could be any use for other tripod beginners as well.
Cheers! Csaba
1.) The best mail which decribes well the everyday life and joy of a microscopist comes first.
You really can't look at a spec sheet and know that it works - it's all trial and error.
2.) About the type of glue to be used:
The best material to use is super glue which is a cyanoacrylate material. There are different types available under various names. They all have slightly different properties and vary in such things as viscosity. The most important point is that the super glue is fresh. Once it is opened, you can use it for a short time, but then need to replace it. Sometimes, the glue from an unopened package can also be bad if it has been on the shelf for a long time. The trouble with using consumer products is that they sometime change without warning.
You should buy it from a place that has a high turnover rate of it so that it is fresh. The IBM guys recommend keeping the stuff unopened in the refrigerator for any length of time. Once it has been opened, you can't use it for very long (a day or two). I have used the Loctite a little longer because it has a very good sealing top. The Loctite product is also available in a "pen" type applicator which seems to have the best bench life of any of the applicators I've used. My tube usually is swiped off the bench long before it goes bad.One of the advantages to the cyanoacrylic cement is it dissolves, in a reasonable time, in acetone.If you were to use a crosslinked epoxy, you'll need to devise a cleaning scheme that will exhaustively remove the epoxy without altering your sample.
Basically what we're looking for is an intermediate viscosity glue (somewhere between maple syrup and water) that bonds in about 30-40 seconds and is very strong. Don't use any type of super glue gel! It doesn't work, you have to use the thin stuff.
In the end we are using a Loctite Prism 460.
Crystalbond 509. This is an acetone soluble glue which melts at 150*C and becomes quite viscous. We use it quite often here at Queens as a temporary glue for ceramics that have to be ground and polished on all four sides. If the bond breaks you simply heat it up and reset the part. This stuff comes in sticks that last for a very long time. Their website is as follows. {http://www.aremco.com/}
I like to glue specimens with epoxy resin because the substance does not harden too fast, and you always have time to find the best location for your sample, so that the latter does not break.
I have tried the "super glue" approach with no luck. Many technicians advocate Lock Tite brand of super glue that many companies sell with the tripod polish kit. I have always utilized Crystalbond adhesive. It is a low melting point (77 C) wax that is dissolved in acetone. If your sample is heat sensitive, super glue is the only other choice I know.
3.)Glueing advices:
--the biggest reason for sections falling off is the cleanliness of the pedastal. It must be cleaned with clean solvents that do not leave any trace of a contamination film on the glass. Check for cleanliness by holding the tool such that light reflects from the surface.
--the suface that you are glueing to should be as rough as possible to obtain a tooth or larger surface area for the glue to adhere to.
--any way you can improve the surface area would be great.
--the same above would be for the specimen
--pressure on the sample while it cures might be important, but the IBM guys just wick the extra stuff away from the sample and don't put a lot of pressure on the sample.
-- ____________________________________________ Csaba Cserhati Univ.of Debrecen / Dept. of Solid State Phys. Hungary tel/fax: 36 52 316073 e-mail: cserhati-at-delfin.klte.hu ____________________________________________
I've followed with interest the discussion on equipment depreciation and service contracts . I've noticed over the years in both metals research and now the semiconductor industry with increasing sophistication/specialisation of equipment running a lab to a given budget seems to mean accepting manufacturers service contracts with fewer 'independent' sources being able or willing to offer maintenance assistance .
My question arises from a recent 'confocal service contract' letter and I wonder does an insurance contract service alternative as offered by CIC exist in the UK ? and if so could anyone let me know where I could get more information ? .
Martyn Harris Device Engineer ESM Ltd , Cardiff Rd Newport , South Wales UK NP10 8YJ .
First, a disclaimer. I am a third party service provider for a variety of equipment, but not confocals. I have worked for a number of manufacturers in the past and been self-employed for around 20 years.
In regards to the first question - service departments in general are getting squirrellier. Contract prices have gone up greatly over the last decade or so while quality and parts stores have gone down. A generalization, granted, but one I'd be happy to back up.
As for the second question - don't get me started again. Given recent problems with the industry in general, I can only suggest that your university carefully study the question and check with several of their customers who have been with them for at least a couple of years insuring similar equipment. That goes for any such provider. I've been a vocal antagonist here of the concept, and from what I have heard from some customers, perhaps rightfully so. Not right, so far, in my feelings regarding the potential long term problems. Rather in the broad approach they have taken. Perhaps in trying to be a jack of all trades, they are a master of none.
To those of you who might be surprised by my abnormally low tone in this posting, please understand that we are getting to a point in time where these organizations are getting quite large in a very short period of time. As with any company or industry, they do have problems that they will not publicize. They also have increasingly large budgets for legal teams that would probably be anxious to root out any libel. They do save many organizations large amounts of money, but you have to ask, at what cost?
On Wednesday, April 04, 2001 10:20 AM, Tom Phillips [SMTP:PhillipsT-at-missouri.edu] wrote: } } } I have two questions concerning service contracts on confocals. Let } me start by saying i have had a confocal for about 8 years and would } never consider going without one. I have a Biorad 2000 going off } warranty and need to make a decision. } } First question: Biorad no longer guarantees a response time - they } now promise to get to you as fast as they can but no longer promise a } 48 or 72 hr response. Have other confocal manufacturers done this } also? } } Second question: My university is pushing replacing service } contracts with "insurance" contracts with a major vendor who then } pays for a service visit from the manufacturer on an hourly basis. } All parts, travel, service repair time, etc are covered at a price } that is typically 75% less than the manufacturer's service contract. } They guarantee the price and coverage for 3 years. Personally I } don't know how they could make money on this deal since we average a } fair number of visits and spare parts (e.g. lasers) in a typical } year. Does anyone have experience with this type of situation with } confocals? The company the University is dealing with is CIC but } there are several other ones out there. } } Thanks for any input. } -- } 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) } }
Allen R. Sampson, Owner Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174 voice 630.513.7093 fax 630.513.7092
I don't know if this is exactly what you are looking for, but Reetz and Coworkers have recently published a high troughput routine for nanaoparticle analysis: Reetz, Manfred T.; Maase, Matthias; Schilling, Tobias; Tesche, Bernd. Computer Image Processing of Transmission Electron Micrograph Pictures as a Fast and Reliable Tool To Analyze the Size of Nanoparticles. J. Phys. Chem. B (2000), 104(37), 8779-8781.
Good Luck,
Andreas
************************************************* Dr. Andreas Taubert Dept. of Materials Science and Engineering 3231 Walnut Street University of Pennsylvania Philadelphia PA 19104-6272 tel: +1 215 898 2700 fax: +1 215 573 2128
Physical Chemistry is everything for which 1/T is linear ... *************************************************
Tina: We have the Duoscan 1200 but the 2500 is also a very nice unit--additional (real) resolution and a high O.D. range plus 14 or 16 bit image depth. The Agfa units also come with a built-in transparency plate (rather than having to add on a separate transparency adaptor). I find that color fidelity is very good with the Duoscan, and Agfa provides both reflective and transparency calibration standards. I am currently scanning in 3x4 TEM negatives at 12 bits (yield is about 26MB per image), and scan time is fairly rapid. There is one option that you might want to consider: the DIImage unit is made for up to 4x5 negatives, and I think (can't remember the last time I read the specs--the neurons aren't firing today) that resolution is in the 2700 dpi range--even for the 4x5 size.
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc.
The opinions expressed are solely my own and do not constitute an endorsement of any vendor or manufacturer. I have no fiduciary interest in either company.
On Wed, 4 Apr 2001 17:22:23 -1000 (HST), Tina Carvalho 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- | | A colleague has asked for recommendations for setting up a digital | darkroom (fun to spend someone else's money!). This person would benefit | from a really good scanner that could deal with prints, large format | negatives (4"x5", 3.25"x4") as well as 35 mm slides. At one time I looked | into an Agfa Duoscan T2500. Do any of you have an opinion about this or | other suitable scanners? | | I know this subject comes up regularly, but I don't feel bad about | introducing it again, since technology evolves so quickly! | | Mahalo, | Tina | | http://www.pbrc.hawaii.edu/bemf/microangela | **************************************************************************** | * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * | * Biological Electron Microscope Facility * (808) 956-6251 * | * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* | **************************************************************************** | |
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc. 900 Rigdebury Road Ridgefield, CT 06877 203-798-5448
_______________________________________________________ Send a cool gift with your E-Card http://www.bluemountain.com/giftcenter/
We have a professor here who has a 1 cm x 2.5cm steel layer about 100 um in thickness on a ceramic substrate. The metal layer was sputter deposited onto the ceramic substrate. The ceramic substrate extends past the metal layer. He needs to get a thickness gradient across the steel layer along the 2.5 cm length. He would like to have one end at about 50 microns in thickness and the other at 2 microns with a gradually decreasing thickness gradient. Steps down would be ok although a smooth transition would be better. We have a laser profilometer to measure anything that we produce.
Does anyone out there have any ideas how to do this? Would a tripod polisher work? I thought about electropolishing and masking off portions at a time but I worry about what will happen at the interface between the ceramic and the metal. Could we alter a dimpler?
Nominal grain size of film is about 10 microns (varies with film speed etc but this is the right order of magnitude). Thus to digitize the film to it's nominal limits your scanner should be able to digitize to better than this spatial dimension.
A simple back of the envelope calculation says a spatial resolution of 10 microns is 2540 - dpi..... and as we all know that must be the optical resolution of the scanner not the interpolated resolution. Scanners at this end are obviously more than you need to digitize photo's and get expensive quickly. Also when you see 2 numbers listed as the scanners resolution, believe only the first number, that is the CCD resolution.
Now add your bit depth. 12 bits is the minimium I would shoot for greyscale image, but if your attempting diffraction work the higher the better (i.e. 14 -16 bits+). For color work obviously multiple the bit depth by 3 one for each primary color (RGB). I've seen a number of 36 bit color scanners but not too many 48 bit ones at } 2540 dpi.
Lastly, dit depth is irrelevant if you don't have a high optical density capabilities otherwise your just digitizing noise. The highest value I believe is an OD of 4.0 but this is for DRUM scanners. Flatbed scanners typically run as low as 2.8, upwards to about 3.4 for the best I've seen in a flatbed.
At 09:40 AM 4/5/01 +0000, Ritchie Sims wrote: } "That", for those who may have missed my first post, being the } problem of how to work out what sort of intermediate lens would be } needed to interface a small cheap CCD video camera (or a webcam) so } that it gives a good image when looking into the existing eyepiece } lens of a given optical microscope (in this case the OM of a JEOL 840 } SEM).
You might try a web search for "afocal coupling". See http://www.photosolve.com/xtendascope.asp for example.
This page, and the message below, are for coupling to the eyepiece of telescopes, but the process should be similar with a microscope eyepiece. If you're looking for "small and cheap", put in a wide field eyepiece and hold the camera close to it and see what happens. I suspect you'll have some cropping of the image due to optical mismatch.
- John
} A friend of mind recently bought a fairly expensive SLR digital still } camera, an Olympus C-2500L, } } http://www.olympusamerica.com/product.asp?c=57&p=16&s=12&product=380 } } and has experimented with afocal photography through both his 8-inch f/6 } Dob and a microscope. All he sees in the camera is a small central disk } of light. He got the following explanation from a post on rec.photo. } digital. } } For cameras with LARGE taking lenses (f 2.8, f2.0) [...] there will } be a problem since their entry pupil ( the area of the lens that lets } the image into the lens) is so large compared to the exit pupil (the } opening on the eyepiece that lets the image out of the microscope) } that a large amount of the image will be lost!! This results in } severe vignetting, and so only a small central spot of image in a } dark field is recorded by the digital camera. } } Does this sound reasonable? The optics of this situation are pretty } mysterious to me, so I can't judge. But it seems like increasing the } focal length of the camera lens (zooming in) should compensate for the } effect of the small exit pupil, and that the problem might be more one } of eye relief (he just can't get the lens close enough to the eyepieces, } which I think are a couple of Plossls and the 25mm SMA that comes with } Celestron Dobs).
And here's the response so far.
} Subject: Re: afocal astrophoto problem: exit pupil? } Date: Wed, 4 Oct 2000 17:52:19 -0400 } From: "Michael A. Covington" {See http://www.CovingtonInnovations.com for address} } Organization: MindSpring Enterprises } Newsgroups: sci.astro.amateur } } } Does this sound reasonable? } } No. The camera lens is *supposed* to have a larger entrance pupil than the } exit pupil of the telescope. When doing afocal photography with an SLR the } entrance pupil of the lens is an inch or more in diameter. } } If there is vignetting, it's probably because the camera is the wrong } distance from the eyepiece -- either too close or too far. } } -- } } Clear skies, } } Michael A. Covington / AI Center / The University of Georgia } Author, ASTROPHOTOGRAPHY FOR THE AMATEUR } http://www.CovingtonInnovations.com/astro {} { } } } Subject: Re: afocal astrophoto problem: exit pupil? } Date: Wed, 4 Oct 2000 14:25:48 -0700 } From: "Bob May" {bobmay-at-nethere.com} } Organization: Posted via Supernews, http://www.supernews.com } Newsgroups: sci.astro.amateur } } Bet that when you look at the viewfinder, you will see an image just } like one that you would see if your eye were at a certain distance } from the EP. If that image looks like the eye is too far away from } the EP then it's a sure thing that the camera's lens is too far away } from the EP. That's how it's all done. The camera is nothing more } than an aritificial eye. } -- } Bob May } Remember that computers do exactly what you tell them to do, not what } you think that you told them! } Bob May } } } Subject: Re: afocal astrophoto problem: exit pupil? } Date: Wed, 04 Oct 2000 19:43:43 GMT } From: "Chuck Olson" {chuckolson01-at-home.com} } Organization: -at-Home Network } Newsgroups: sci.astro.amateur } } Yes, it is critical that the eyepiece and camera lens be somewhat } physically compatible with each other. The eyepiece must put the } exit pupil about in the plane of the camera iris opening, which } in most instances requires the eyepiece to be virtually in } contact with the camera lens front element. For instance, the } Nikon Coolpix 950 and 990 have relatively small lens fronts and a } nice 28mm (I think) thread that adapts readily to the T-thread } that is often used in astrophotography. As a result, the CP950 } easily looks through 17mm , 26mm, or 32mm Plossl eyepieces. Even } there, as you point out, the camera needs to be operating at the } tele end of its zoom range to fill the rectangular frame, rather } than showing a small, circular, fuzzy-edged, wide-angle field. } } I'm not sure what the C-2500L looks like, but it may have a } physically larger lens that has its iris deep behind the front } surface. This might require a very long focus eyepiece, like a } 40mm Plossl, conceivably, or one with even greater eye relief, to } accomplish the optical hook-up more favorably, and may limit the } operation of the overall system to somewhat lower magnifications. } Oh, once you have a compatible eyepiece, then you can use Barlow } lenses to get back needed magnification for your desired image } scale. The only probmen there is the setup gets pretty long as } these lenses are stacked up, and stability may suffer. } } The Nikon, as mentioned, has been found by many to be almost } ideal for afocal photography of the moon and planets. } } Chuck }
************************************************************************* {/bigger} {/bold} Interest in the sophisticated fluorescence imaging techniques of Fluorescence Resonance Energy Transfer (FRET) and Fluorescent Lifetime Imaging Microscopy (FLIM) amongst the biological research community has grown in recent years. FRET imaging provides a tool to solve complex structural associations at resolution limits beyond conventional optical imaging. FLIM allows the measurement of FRET without the significant problems associated with intensity based FRET measurement, as well as faster, more accurate and quantitative measurement of cell physiology. These techniques are also being implemented in high-throughput screening regimes for drug discovery. Invited lectures by a distinguished group of scientists will concentrate on new technical developments in these areas and demonstrate successful application of these techniques in biological and industrial settings.
A poster session has been organized for June 9th so that registrants may present their experiences with FRET and FLIM.
We anticipate that this will be a most enjoyable as well as intellectually stimulating symposium.
************************************************************************* {/bigger} {/bold} Interest in the sophisticated fluorescence imaging techniques of Fluorescence Resonance Energy Transfer (FRET) and Fluorescent Lifetime Imaging Microscopy (FLIM) amongst the biological research community has grown in recent years. FRET imaging provides a tool to solve complex structural associations at resolution limits beyond conventional optical imaging. FLIM allows the measurement of FRET without the significant problems associated with intensity based FRET measurement, as well as faster, more accurate and quantitative measurement of cell physiology. These techniques are also being implemented in high-throughput screening regimes for drug discovery. Invited lectures by a distinguished group of scientists will concentrate on new technical developments in these areas and demonstrate successful application of these techniques in biological and industrial settings.
A poster session has been organized for June 9th so that registrants may present their experiences with FRET and FLIM.
We anticipate that this will be a most enjoyable as well as intellectually stimulating symposium.
} Dear colleagues, I would like to buy a digital camera } for my Axiolab Zeiss microscope. } Unfortunately I am a bit confused in the amount of } available data. I would need a digital camera of the } resolution that matches the quality of film cameras } in order I need not scan photos or negatives. } ...
To give you an idea of what you are asking: For comparable resolution, the camera would need deliver more than 6M pixels ... and there is also the question of a digital camera capturing the gamut of color capable of film. For example, the camera you mention, which is aimed at consumers, probably delivers a gamut aimed at the "sRGB" color space. Only a film scanner can capture a color gamut comparable to "Ektaspace RGB". Still, your camera is likely to do a very good job if properly adapted to the microscope. You will need a 1X C-mount adapter for the microscope head, and an adapter for mounting the camera on the C-mount. These are readily obtained for Nikon Coolpix cameras, possibly yours too.
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/
Hello Tina, I have the scanner you are looking at & like it a lot. To be quite honest I do not find that I need to exploit it'll full capably. If I were in the market again, looking at newer technology I would be interested in a faster scanner of similar quality. Yes I want my cake & to eat it too :). I'll give you this analogy. If I have 10 negatives I will franchise my time, that is let things scan while I hang out in the office doing other things. If I have 20 negatives, I'll probably goto the darkroom to make photos. It is quicker & paper is cheaper. BTW I have an Epson 870 inkjet that produces nice quality images... cost is down to $180 US, (now the Epson 880)....no financial interest in these companies.
Oh yea, get the fastest computer you can afford.
good luck, Bruce Brinson Rice U.
Tina Carvalho 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- } } A colleague has asked for recommendations for setting up a digital } darkroom (fun to spend someone else's money!). This person would benefit } from a really good scanner that could deal with prints, large format } negatives (4"x5", 3.25"x4") as well as 35 mm slides. At one time I looked } into an Agfa Duoscan T2500. Do any of you have an opinion about this or } other suitable scanners? } } I know this subject comes up regularly, but I don't feel bad about } introducing it again, since technology evolves so quickly! } } Mahalo, } Tina } } http://www.pbrc.hawaii.edu/bemf/microangela } **************************************************************************** } * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * } * Biological Electron Microscope Facility * (808) 956-6251 * } * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* } ****************************************************************************
Tina, Have your colleague check out the Imacon Flextight Precision II scanner. The optical resolution is 5760 dpi for slide-sized objects; I believe it drops to 4800 dpi for objects the size of her larger negatives. The scanner collects 14 bits of usable data per channel, which can be exported as a two bytes per channel, and has a dynamic range of 3.9 OD units (4.1 OD max). The machine is also very fast. The URL is:
At 05:22 PM 4/4/2001 -1000, Tina Carvalho wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Tina,
I have the Duoscan and a Nikon slide scanner. The Duoscan can scan slides on the special tray feature but side by side comparisons of the Duoscan and Nikon show that the Nikon scan is much better. For the larger negs we had a special tray made for the Duoscan and we scan in our EM negs. The Nikon has gotten much cheaper and an excellent scanner can be had for $700 with Digital ICE, something you want.
Get two scanners.
Rick A. Harris, Director Microscopy and Imaging Facility Section of Molecular and Cellular Biology 1241 Life Sciences Addition University of California Davis, CA 530 752 2914 530 754 7536 fax http://katie.ucdavis.edu raharris-at-ucdavis.edu
I forgot to mention John that we do not use formar coated grids. For better assessment of the renal biopsy we do not mince the sample, either. Please see our Web page for more details. {pathology.lsuhsc.edu/Pathist/dx_home.htlm} click on M.diagnostic service and then on renal biopsy. Our diagnostic em lab continues to send poloroid HRLM pictures and TEM B&W contact prints with each report.
Lehigh University seeks an Electron Microscopy Technician to perform duties in support of the Microscopy Center of the Materials Science and Engineering Department. The person appointed will work with other technical staff to instruct students in the operation of microscopes and other equipment; maintain and repair instruments; carry out upkeep of the lab; support research professors and students; analyze samples; give tours and demonstrations; maintain a safe environment and perform other assigned duties. A bachelor's degree in physical science and/or 4+ years related work experience is required. Candidates should be familiar with electron microscopes, mechanical and electronic equipment, vacuum systems, computers (PC and/or Mac) and EDS/WDS systems. Experience with a microprobe would be especially valuable. Good communication and interpersonal skills are essential.
Lehigh University offers excellent benefits including medical, vision and tuition. Interested candidates should forward their resume to Jennifer Mohney, Human Resources, 428 Brodhead Avenue, Lehigh University, Bethlehem PA 18015. EEO/AA
-- .......... Alwyn Eades Department of Materials Science and Engineering Lehigh University 5 East Packer Avenue Bethlehem Pennsylvania 18015-3195 Phone 610 758 4231 Fax 610 758 4244 jae5-at-lehigh.edu
One of our students is trying to stain the DNA of osteoblasts using bisBenzamid, Hoechst no. 33258 trihydrochloride. She is finding conflicting information about the concentration to use and the lethal dose.
I would appreciate it if someone can provide or point us to a protocol they have used successfully. We are primarily a materials lab and do not have a lot of biological reference material at hand.
One of our students is trying to stain the DNA of osteoblasts using bisBenzamid, Hoechst no. 33258 trihydrochloride. She is finding conflicting information about the concentration to use and the lethal dose.
I would appreciate it if someone can provide or point us to a protocol they have used successfully. We are primarily a materials lab and do not have a lot of biological reference material at hand.
I was initially hesitant to introduce a subject that gets periodically posted here, but received so many enthusiastic messages about print and negative scanners that I thought I'd continue the thought. I will be happy to summarize the responses and throw in some opinions of my own. As more and more of us convert to "digital darkrooms" we can share more of our experiences with hardware and software.
I am helping a former traditional photographic media user/computerphobe set up digital imaging capabilities since her university/museum department is closing their darkroom facilities and reassigning personnel (sigh). She has what appears to be a decent budget (until I started pricing the good stuff!). I am proposing she get a fast (733MHz) G4 Mac with maximum (1.5GB) RAM, and she saw and fell in love with the Apple 22" cinema display (as did I when I saw it in person). People seem to like the Agfa T2500 scanner for prints and negatives. A moderate color printer, since she has access to other really good printers in her department. A Nikon Coolpix 990 digital camera for on- and off-microscope. Photoshop 6.0, for which I'll train her. Corel Draw for vector graphics?
Additions, subtractions and comments will be welcome. I'll summarize after a reasonable amount of time and we'll see if there is a consensus on the ideal digital darkroom!
Mahalo, Tina
http://www.pbrc.hawaii.edu/bemf/microangela
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
How does your institution handle short term storage in the lab of waste osmium tetroxide. Does it need to be in a fume hood? We keep it in closed glass containers mixed with vegetable oil. Does anyone have a reference regarding whether or not it needs to be stored in a hood? Thank you for any responses.
John, I forgot to mention that we do not use formar coated grids. For better assessment of the renal biopsy we do not mince the sample, either. Please see our Web page for more details. {pathology.lsuhsc.edu/Pathist/dx_home.htlm} click on M.diagnostic service and then on renal biopsy. Our diagnostic em lab continues to send poloroid HRLM pictures and TEM B&W contact prints with each report.
I am a laboratory instructor at Brookdale Community College in Lincroft, NJ who operates a Zeiss 9C TEM which was donated to the college. We are still in the process of being able to make our own grids. Does anyone have some grids that they would be willing to sell or donate to the College? Grids of anything would be greatly appreciated, but basic cellular structures is really what we are looking for as we are a community college and only have first and second year students. Thanks in advance for your help.
I used Agfa DuoScan HiD earlier and I try to get it here as well. I like that machine a lot. It's optical resolution is 1000x2000 Dynamic range is 3.7D, which would help scanning DP's. If you want more info you can have a look at:
http://www.agfa.com/scanners/duoscan_HiD.html
Printing is another task you can buy things from AGFA as well. Their photoprinter is just excellent, but a bit expensive. I have tried nice HP injet printers with great success.
Cheers! Csaba
-- ____________________________________________ Csaba Cserhati Univ.of Debrecen / Dept. of Solid State Phys. Hungary tel/fax: 36 52 316073 e-mail: cserhati-at-delfin.klte.hu ____________________________________________
I used Agfa DuoScan HiD earlier and I try to get it here as well. I like that machine a lot. It's optical resolution is 1000x2000 Dynamic range is 3.7D, which would help scanning DP's. If you want more info you can have a look at:
http://www.agfa.com/scanners/duoscan_HiD.html
Printing is another task you can buy things from AGFA as well. Their photoprinter is just excellent, but a bit expensive. I have tried nice HP injet printers with great success.
Cheers! Csaba
-- ____________________________________________ Csaba Cserhati Univ.of Debrecen / Dept. of Solid State Phys. Hungary tel/fax: 36 52 316073 e-mail: cserhati-at-delfin.klte.hu ____________________________________________
I used Agfa DuoScan HiD earlier and I try to get it here as well. I like that machine a lot. It's optical resolution is 1000x2000 Dynamic range is 3.7D, which would help scanning DP's. If you want more info you can have a look at:
http://www.agfa.com/scanners/duoscan_HiD.html
Printing is another task you can buy things from AGFA as well. Their photoprinter is just excellent, but a bit expensive. I have tried nice HP injet printers with great success.
Cheers! Csaba
-- ____________________________________________ Csaba Cserhati Univ.of Debrecen / Dept. of Solid State Phys. Hungary tel/fax: 36 52 316073 e-mail: cserhati-at-delfin.klte.hu ____________________________________________
Based on historical precedent we keep used osmium tetroxide in the fridge in a "Kilner" jar (for pickled fruit and veg.), which has a rubber seal.
BTW what ratio of vegetable oil to osmium solution do you use?
Dave
On Thu, 5 Apr 2001 20:30:30 -0500 Michele von Turkovich {mvonturk-at-zoo.uvm.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 } -----------------------------------------------------------------------. } } } How does your institution handle short term storage in the lab of waste } osmium tetroxide. Does it need to be in a fume hood? We keep it in closed } glass containers mixed with vegetable oil. Does anyone have a reference } regarding whether or not it needs to be stored in a hood? Thank you for any } responses. } } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
In respose to Tina's post, I have not seen any mention on the list of the scanner I purchased a few weeks ago, the Epson Expression 1640XL. It has 1600dpi optical resolution (scans at a hardware resolution of 1600x3200 dpi) 42 bit color (14 bit gray) and Dmax of 3.6. It is large format, and the transparency adapter comes with a range of negative holders. Has SCSI or USB interfaces with firewire as an optional extra (I use USB on a Win 2000 system). Of course, you pay for what you get - it isn't cheap.
We are only just beginning to learn how best to use all the resolution and bit depth we now have, but I and my users love it!
This is not a comparison, of course (I haven't used the other models) but just to say we are happy with what we have.
Tony.
} } Hi, All- } } A colleague has asked for recommendations for setting up a digital } darkroom (fun to spend someone else's money!). This person would benefit } from a really good scanner that could deal with prints, large format } negatives (4"x5", 3.25"x4") as well as 35 mm slides. At one time I looked } into an Agfa Duoscan T2500. Do any of you have an opinion about this or } other suitable scanners? } } I know this subject comes up regularly, but I don't feel bad about } introducing it again, since technology evolves so quickly! } } Mahalo, } Tina } } http://www.pbrc.hawaii.edu/bemf/microangela } **************************************************************************** } * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * } * Biological Electron Microscope Facility * (808) 956-6251 * } * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* } **************************************************************************** }
Hysterical precedents aside. The idea of the oil is to absorb all tetroxide vapours and render the substance harmless. Metallic Os is essentially non-toxic, so the treated tetroxide waste should smell of whatever vegetable oil and not emit any of the musky smell emitted by osmium tetroxide. Neither should the treated material be regarded as hazardous waste - but I expect that no safety officer would have the courage to make that declaration. 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 ABN: 99 724 136 560 www.proscitech.com
On Friday, April 06, 2001 6:52 PM, Patton, David [SMTP:David.Patton-at-uwe.ac.uk] wrote: } } } } Based on historical precedent we keep used osmium tetroxide } in the fridge in a "Kilner" jar (for pickled fruit and } veg.), which has a rubber seal. } } BTW what ratio of vegetable oil to osmium solution do you } use? } } Dave } } } On Thu, 5 Apr 2001 20:30:30 -0500 Michele von Turkovich } {mvonturk-at-zoo.uvm.edu} wrote: } } } } } } } How does your institution handle short term storage in the lab of waste } } osmium tetroxide. Does it need to be in a fume hood? We keep it in closed } } glass containers mixed with vegetable oil. Does anyone have a reference } } regarding whether or not it needs to be stored in a hood? Thank you for any } } responses. } } } } } } } } ---------------------------------------- } Patton, David } Email: David.Patton-at-uwe.ac.uk } "University of the West of England" } } }
between membranes in all cellular components that have two or more membrances and
also seeking "volume and size" of all known subcellular organelles with or without membranes. also seeking " other physical measurement parameters" of cellular components by cell type.. by cell age.. etc. thanks.
Computer Aided Cell and Molecular Biology (CACMB), not medicine, will find the cure for cancer and other diseases. There will always be a need for the trained clinician (MD/RN) but, advanced diagnostic and treatment option selection has become gene based, has moved from the physician's practice to the computerized cell and molecular biology laboratory, and appropriate treatment options should now be based on the personal biology of the patient.
Adobe Illustrator or Freehand for vector graphics. Corel Draw is less common and has a proprietary file format that has caused me troubles preparing articles for Microscopy Today. Corel can save in other formats, but people have to use that option. Also, Adobe has educational pricing, and special package prices that bundle full versions of Photoshop and Illustrator.
Phil
} Hi, again- } } I was initially hesitant to introduce a subject that gets periodically } posted here, but received so many enthusiastic messages about print and } negative scanners that I thought I'd continue the thought. I will be happy } to summarize the responses and throw in some opinions of my own. As more } and more of us convert to "digital darkrooms" we can share more of our } experiences with hardware and software. } } I am helping a former traditional photographic media user/computerphobe } set up digital imaging capabilities since her university/museum department } is closing their darkroom facilities and reassigning personnel (sigh). She } has what appears to be a decent budget (until I started pricing the good } stuff!). I am proposing she get a fast (733MHz) G4 Mac with maximum } (1.5GB) RAM, and she saw and fell in love with the Apple 22" cinema } display (as did I when I saw it in person). People seem to like } the Agfa T2500 scanner for prints and negatives. A moderate color printer, } since she has access to other really good printers in her department. A } Nikon Coolpix 990 digital camera for on- and off-microscope. Photoshop } 6.0, for which I'll train her. Corel Draw for vector graphics? } } Additions, subtractions and comments will be welcome. I'll summarize after } a reasonable amount of time and we'll see if there is a consensus on the } ideal digital darkroom! } } Mahalo, } Tina } } http://www.pbrc.hawaii.edu/bemf/microangela } } **************************************************************************** } * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * } * Biological Electron Microscope Facility * (808) 956-6251 * } * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* } ****************************************************************************
-- }}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{ Philip Oshel Supervisor, AMFSC and BBPIC microscopy facilities Department of Animal Sciences University of Wisconsin 1675 Observatory Drive Madison, WI 53706 - 1284 voice: (608) 263-4162 fax: (608) 262-5157 (dept. fax)
We keep used osmium in small sealed bottles in the hood. Our environmentals services office would rather have **undiluted/unmixed** (no oil) waste for them to neutralize. They have no way of knowing whether all the osmium has been denatured by attachment to oil and would have to treat the whole bottle as osmium waste making for more volume to have to dispose of. Check with your hazardous waste office.
Sara E. Miller, Ph. D. P. O. Box 3712 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-3265
Email: inikolak-at-mred.tuc.gr or eisodos-at-otenet. Name: Irene Nikolakaki
Organization: Technical University of Crete
Education: Graduate College
Location: Chanea, Crete,Greece
Question: Dear sir/madam,
I am a postgraduate student at the Technical University of Crete and I seek information (as detailed as possible) on how enumeration of airborne microorganisms collected on Nuclepore filters is done with the use of epi-fluorescence microscopy.I would be grateful if you could provide me with this infromation or any kind of help as to where I can find it.
I both run the microscope labs here and am a researcher in materials engineering.
First I want to say that I strongly support the use of the digital laboratory. While we still occasionally use film for our highest quality requirements, in general we are fully digital. The use of digital cameras has really expanded our undergraduate teaching laboratories and has sped up our research.
I have found one "dark-side" to a digital imaging laboratory as a lab manager. As the lab manager, I have found that keeping a digital laboratory up-to-date is much more expensive than the film laboratory. When we were only film, we had to repair the film cartridges for our Polaroid PN film (it takes about five minutes) and have the microscopes cleaned about once a year at a cost of about $1k.
The digital lab. is much more expensive time and repair wise. Because we crunch our computers with our image size and storage, it takes more of my time to keep stuff going. All our computers are networked and in addition to work, the students tend to junk up the computers with downloads etc which stops them from working for the image processing work. This requires continual monitoring on my part (in spite of rules against using them for these applications!) In addition, keeping computers that will run the data is expensive. I buy pretty much the best out there, but somehow upgrades are still inevitable. I also have to supply print cartridges,etc. Researchers always supplied their own film and dark room supplies. In addition, I've had to have our cameras repaired numerous times. The cost was high (at least $500) and they stayed gone for up to a month. Finally, some of my cameras are about 3 years old. I can see a degredation in the image quality from when they were purchased. The cameras are much noisier. I see a future of regular replacement of my cameras in addition to the computer upgrades. So while the cost to the researchers is lower (which helps me as a researcher), the cost to the lab itself is higher (which hurts me as a lab manager). I'm working on setting up a fee schedule for this equipment but REALLY hate to have to do it. All of you who do this in a university know how painful it is!
Regarding the camera purchase-in addition to considering the camera resolution and cost, I think you should consider the image transfer. I recommend considering a camera with immediate transfer of the image to the microscope if you have numerous inexperienced users. Being able to focus on the screen is extremely helpful. The image transfer time is also important if you have many images to capture. We do image analysis on numerous images and some of the cameras have about a 30 second transfer time for decent resolution. This would be unbearable for the number of images we collect. I'm not sure how the Nikon Coolpix works but this should be considered by anyone that is purchasing a digital camera.
Good luck!
Robin Griffin UAB
-----Original Message----- } From: Tina Carvalho [mailto:tina-at-pbrc.hawaii.edu] Sent: Thursday, April 05, 2001 6:09 PM To: Microscopy Listserver
Hi, again-
I was initially hesitant to introduce a subject that gets periodically posted here, but received so many enthusiastic messages about print and negative scanners that I thought I'd continue the thought. I will be happy to summarize the responses and throw in some opinions of my own. As more and more of us convert to "digital darkrooms" we can share more of our experiences with hardware and software.
I am helping a former traditional photographic media user/computerphobe set up digital imaging capabilities since her university/museum department is closing their darkroom facilities and reassigning personnel (sigh). She has what appears to be a decent budget (until I started pricing the good stuff!). I am proposing she get a fast (733MHz) G4 Mac with maximum (1.5GB) RAM, and she saw and fell in love with the Apple 22" cinema display (as did I when I saw it in person). People seem to like the Agfa T2500 scanner for prints and negatives. A moderate color printer, since she has access to other really good printers in her department. A Nikon Coolpix 990 digital camera for on- and off-microscope. Photoshop 6.0, for which I'll train her. Corel Draw for vector graphics?
Additions, subtractions and comments will be welcome. I'll summarize after a reasonable amount of time and we'll see if there is a consensus on the ideal digital darkroom!
Mahalo, Tina
http://www.pbrc.hawaii.edu/bemf/microangela
**************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu *
* Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf*
Hello, On the September/October issue of Microscopy and Microanalysis, they have a cover picture of a yeast SEM image collected by David Scharf. I was wondering if anyone knew the details of how the sample was prepared. It looks as though the sample was processed directly from the medium it was growing on. I am used to processing bacteria and yeast by filtering through membrane filters, or depositing on polylysine treated glass/silica. I was wondering if there was something different done for this sample?
\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\ 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
I am in need of some form of a conductive film coater for SEM sample preparation. I have found a Tousimis Research Corporation Samsputter-2a sputter coater, though it may not be working properly.
Until now, I have been considering constructing my own sputter coater from microwave oven parts and the like, but I have not yet found any books or other resources with clear enough information to get my confidence to the point that I feel that I can do it. I have an understanding of what is needed electrically, but I need something that fills in some details like "Be sure that the target to sample distance can be adjusted between X and Y. You need to get a QRZ type needle valve to admit the argon", etc.
Does anyone have experience with the Tousimis Samsputter 2a? From what I can see, it is an extremely simple unit. Is it something that would be worth repairing? Would it best be used to scavenge parts for a home-built unit?
Since I have breeched the question of home-built sputter coaters, does anyone know of any resources that describe the dos and donts of building one? I know that there are a number of vendors on the listserv. What are the critical features of your units that I just can't get in a home-built unit and just can't live without?
To put this all in perspective, I have an old SEM with 100 Angstrom resolution at best. I don't need flawless, grainless deposition. I have close to zero budget. Even a used sputter coater at market prices is far too expensive. Either I make something like this work, or I learn to work with uncoated samples (which will be difficult, since I am interested in looking at clays - small particle size with next to zero conductivity).
I suggest to use SEM (with calibrated magnification) if you could electroplate your sample with layer of Ni 0.1-0.3 mm thick. Sorry, I do not have a protocol for plating with Ni now, but I did it many times and it is pretty easy.
After coating with Ni you can cut your sample, mount it in epoxy or thermoset and prepare usual cross section. Ni would save the edge of steel layer and measurements will be very simple.
If surface of you samples is flat, you can cut them (better with diamond saw), mount them in epoxy with a steel strip (instead of Ni) attached to steel layer and polish. It is less dependable but still not bad method.
Vladimir Dusevich
-----Original Message----- } From: "rgriffin-at-eng.uab.edu"-at-sparc5.microscopy.com [mailto:"rgriffin-at-eng.uab.edu"-at-sparc5.microscopy.com] Sent: Thursday, April 05, 2001 8:25 AM To: microscopy-at-sparc5.microscopy.com Cc: hban-at-eng.uab.edu
We have a professor here who has a 1 cm x 2.5cm steel layer about 100 um in thickness on a ceramic substrate. The metal layer was sputter deposited onto the ceramic substrate. The ceramic substrate extends past the metal layer. He needs to get a thickness gradient across the steel layer along the 2.5 cm length. He would like to have one end at about 50 microns in thickness and the other at 2 microns with a gradually decreasing thickness gradient. Steps down would be ok although a smooth transition would be better. We have a laser profilometer to measure anything that we produce.
Does anyone out there have any ideas how to do this? Would a tripod polisher work? I thought about electropolishing and masking off portions at a time but I worry about what will happen at the interface between the ceramic and the metal. Could we alter a dimpler?
Osmium waste is toxic and hazardous due to its reactivity and heavy metal characteristics. One of the "urban legends" that abounds is that Osmium mixed with oil is "neutralized" and made safe. It is true that much of it is reduced to metallic Osmium. However in a bizarre incident it was shown in our lab years ago that reduced Osmium is very reactive with strong oxidizers. In the incident some reduced Osmium was accidentally spilled in a sink which had a small amount of Hydrogen Peroxide in the drain. The ensuing yellow gas cloud of newly formed Osmium Tetroxide from the exothermic reaction was very toxic!!!! I mention this only to confirm that Sara Miller is right again as usual and that mixing Osmium waste with other compounds does not make it safe and does increase the volume.
Hi All, I've been tearing hair out all day trying to find a recipe for GMA. In the past, I have used either JB-4 or Historesin (or Technovit) embedding kits, which have clear directions for embedding tissue and polymerizing these resins without UV. I recently ordered a straight GMA embedding kit (because it was much cheaper), and received not one bit of direction as to how I should combine the three components. I did find a protocol on EMS' website for UV polymerization, but would much prefer to use the non-UV method, since it allows tissue to be better oriented. Before I start experimenting, I thought I'd see if anyone out there can help. Thanks again for your help. Bald in Iowa, Kristen Kristen A. Lennon, Ph.D. Department of Plant Pathology 351 Bessey Hall Iowa State University Ames, IA 50011 515-294-8854 kalen-at-iastate.edu
Osmium metal is sometimes reported not to react with air, but other sources report slow reactivity of the metal with air to produce OsO4. It is interesting to note that the name osmium is derived from the Greek "osme" meaning smell, referring to the odour of the metal resulting from osmium tetroxide production at its surface. Presumably neither the finely divided metal nor the dioxide (osmium black) can be trusted not to oxidise to OsO4 in air. What makes osmium tetroxide especially hazardous is the fact that it is very volatile. Unlike OsO4 neither osmium metal nor osmium dioxide are volatile, and provided oxygen can be prevented from reaching them they are therefore relatively innocuous. Surrounding them in oil is probably therefore a good strategy, provided the mixture is still treated as toxic waste.
Dr. Chris Jeffree Inveresk Cottage 26, Carberry Road Inveresk Musselburgh Midlothian EH21 8PR Tel: +44 131 665 6062 FAX +44 131 653 6248 Mobile 07710 585 401
I think that I have missed part of this message string somewhere, but it sounds like you are starting with a uniform thickness steel layer that has been deposited on a ceramic substrate. I will then make the assumption that you are attempting to alter the thickness of the steel layer across the length of the sample to produce a 'wedge'. I am not sure that this will help, but you could try the wedge polishing technique used for TEM sample preparation, or bevel polishing. I have had success bevel polishing IC's, but not to the degree of accuracy that you require. I know that South Bay technologies makes a pretty good tripod with micrometer levels at all three corners that may help you out. The biggest problem will be in setting up to ensure a good gradient. You should probably try it a few times with 'dummy' samples to get the feel for how aggressive your polish angle is. I hope this helps. Nick Aitken
-----Original Message----- } From: Dusevich, Vladimir [mailto:DusevichV-at-umkc.edu] Sent: Friday, April 06, 2001 11:37 AM To: microscopy-at-sparc5.microscopy.com Cc: hban-at-eng.uab.edu
I suggest to use SEM (with calibrated magnification) if you could electroplate your sample with layer of Ni 0.1-0.3 mm thick. Sorry, I do not have a protocol for plating with Ni now, but I did it many times and it is pretty easy.
After coating with Ni you can cut your sample, mount it in epoxy or thermoset and prepare usual cross section. Ni would save the edge of steel layer and measurements will be very simple.
If surface of you samples is flat, you can cut them (better with diamond saw), mount them in epoxy with a steel strip (instead of Ni) attached to steel layer and polish. It is less dependable but still not bad method.
Vladimir Dusevich
-----Original Message----- } From: "rgriffin-at-eng.uab.edu"-at-sparc5.microscopy.com [mailto:"rgriffin-at-eng.uab.edu"-at-sparc5.microscopy.com] Sent: Thursday, April 05, 2001 8:25 AM To: microscopy-at-sparc5.microscopy.com Cc: hban-at-eng.uab.edu
We have a professor here who has a 1 cm x 2.5cm steel layer about 100 um in thickness on a ceramic substrate. The metal layer was sputter deposited onto the ceramic substrate. The ceramic substrate extends past the metal layer. He needs to get a thickness gradient across the steel layer along the 2.5 cm length. He would like to have one end at about 50 microns in thickness and the other at 2 microns with a gradually decreasing thickness gradient. Steps down would be ok although a smooth transition would be better. We have a laser profilometer to measure anything that we produce.
Does anyone out there have any ideas how to do this? Would a tripod polisher work? I thought about electropolishing and masking off portions at a time but I worry about what will happen at the interface between the ceramic and the metal. Could we alter a dimpler?
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Jim Darley wrote: ================================================== Hysterical precedents aside. The idea of the oil is to absorb all tetroxide vapours and render the substance harmless. Metallic Os is essentially non-toxic, so the treated tetroxide waste should smell of whatever vegetable oil and not emit any of the musky smell emitted by osmium tetroxide. Neither should the treated material be regarded as hazardous waste - but I expect that no safety officer would have the courage to make that declaration. ================================================== Jim might be right but there are other points of view on this:
1] I have been led to believe that the vegetable oil reduction of remaining unreduced tetroxide reduces things down to the dioxide, not the metal. The dioxide is a black colloidal solid (the metal is a lighter "bluish gray"). Osmium dioxide is itself relatively innocuous, for example, it is not "regulated" as a dangerous good when shipped, either domestically under US DOT rules or internationally under IATA rules.
2] Osmium is a non-renewable resource and as such, we should all be looking for ways to recycle such materials as opposed to taking them to landfills or in the case of osmium, incineration. I am no environmental "activist" myself, but I think we should all be thinking about such things. The possibility of recycling something however, is intimately connected with the economics of recycling vs. the cost of the purchase of new virgin material. And when the "used" osmium tetroxide containing aqueous liquid is "neutralized" in vegetable oil, for those involved in precious metals recycling, this act essentially kills the economics of recycling.
3] We have in beta testing stage right now an "osmium recycling kit". We are looking for a limited number of laboratories (for now, just in the USA) to participate in our beta testing of this kit. If you are interested in participating in this test, let me know off-line and I will send you the details.
4] In the mean time, I would offer the following advice. Consider using one of the other methods described on this listserver in the past, such as the ones involving KOH as a reducing agent. The reduced material in this state can be recycled economically, but only in large quantities. No one laboratory, in our opinion, could generate enough such material over a reasonable period of time, to make recycling and refining, even of the material is in this state, economical.
5] The one thing you don't ever want to do, at least in the USA, is to declare this as any kind of a "hazardous waste". Once something has been declared to be a hazardous waste, that designation can never (if my understanding of regulations is correct) be reversed, and it forever has to be treated as a waste, and translated, that means it is destined for eventual disposal by either landfill or incineration. I want to be very careful here, it is complicated, this is true so long as we are talking about a RCRA hazardous waste, that is, it meets a listing or characteristic definition. One environmental experts tells me the following: "The reason OsO4 (and OsO2, for that matter) are not regulated as hazardous has nothing to do with their human toxicity (or lack of); it is because osmium, from a regulatory standpoint, is not considered toxic to the environment." He also says it is OK to designate something as a hazardous waste with the intent to recycle it; it simply must be managed as a hazardous waste while it is on-site.
So these containers that are holding reduced material (from the tetroxide) for recycling must be labeled properly, and that would mean something like "osmium dioxide for recycling". I am getting into an area that is not black and white defined, and probably varies from institution to institution in the US, not to mention the variation from country to country. But the important thing is that from a regulatory point of view, what that label says ends up determining the possibilities for the ultimate fate of its contents.
We are striving to find a way to help people transfer, on their environmental "accounting sheets", a material from the column saying "incineration" or "landfill", to the "recylcing" column. And for those who worry about such things, from a legal liability standpoint, there is general recognition that if the material is recycled, there is far less legal liability associated with its disposal than if it is incinerated or sent to landfill. And at the same time, recycling keeps this most valuable nonrenewable resource in the stream of commerce and available for future generations.
Disclaimer: SPI Supplies has developed a kit for recycling osmium. It is not yet commercially available, but is in beta test stage. We also have the obviously ulterior motive of making sure there is osmium tetroxide available for future generations of EM users, otherwise there would be no place for firms like SPI Supplies.......
Chuck
PS: Please remember that we are nearly 100% paperless and we would ask that any reply to this message be by way of the "reply" feature on your software, so that the entire string of correspondence can come back to us and all be in one place.
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Different people approach such subjects from rather different angles and they may all be right, but considering their own realities.
A couple of submissions approached the subject from an "absolute safety" point of view. I don't subscribe to that because the amount of tetroxide emitted from gram quantities of metal and some dioxide when surrounded by the tetroxide absorbing oil, would be miniscule. In my opinion, Os waste in vegetable oil is not a substantial hazard in the lab or when dumped. For many, particularly in the USA this is irrelevant, because of strict legal obligations.
There is also an environmental angle. Unfortunately, greatest safety regardless of cost, is at odds with good environmental practice. If a lot of material and energy is used to dispose of a low hazard material, then the total environmental cost may be very high when compared with an "acceptable risk" solution. We should not confuse the ever-increasing demands for greatest safety with "environmentally friendly" practices: they are not synonymous, but often mutually exclusive.
I was pleased to read some of Chuck's submission. Whatever the motivation, recycling of a limited resource is commendable. This has been tried before by turning the waste material back into the tetroxide. I understand that few people still practice the regeneration of osmium. One reason is that the actual fixative solution is poorly defined after reclamation from diverse fixation vehicles.
Chuck's idea of turning the material into metallic osmium is possible. Using precision electrolysis, for instance with SS electrodes at ~400volts, with the plate size determining amperage, the recovered metal could be 99.9% pure. The refiner, however, would charge for assaying and refining and this means another business would need to collect the osmium, resulting in double shipping and double mark-ups. Considering the few grams recoverable, instrument cost and maintenance, this would be a marginal business, but one that the larger labs certainly should consider. 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 ABN: 99 724 136 560 www.proscitech.com
On Monday, April 09, 2001 4:45 AM, Garber, Charles A. [SMTP:cgarber-at-2spi.com] wrote: } } } -- [ From: Garber, Charles A. * EMC.Ver #3.1 ] -- } } Jim Darley wrote: } ================================================== } Hysterical precedents aside. The idea of the oil is to absorb all tetroxide } vapours and render the substance harmless. Metallic Os is essentially } non-toxic, so the treated tetroxide waste should smell of whatever } vegetable } oil and not emit any of the musky smell emitted by osmium tetroxide. } Neither should the treated material be regarded as hazardous waste - but I } expect that no safety officer would have the courage to make that } declaration. } ================================================== } Jim might be right but there are other points of view on this: } } 1] I have been led to believe that the vegetable oil reduction of remaining } unreduced tetroxide reduces things down to the dioxide, not the metal. The } dioxide is a black colloidal solid (the metal is a lighter "bluish gray"). } Osmium dioxide is itself relatively innocuous, for example, it is not } "regulated" as a dangerous good when shipped, either domestically under US } DOT rules or internationally under IATA rules. } } 2] Osmium is a non-renewable resource and as such, we should all be looking } for ways to recycle such materials as opposed to taking them to landfills or } in the case of osmium, incineration. I am no environmental "activist" } myself, but I think we should all be thinking about such things. The } possibility of recycling something however, is intimately connected with the } economics of recycling vs. the cost of the purchase of new virgin material. } And when the "used" osmium tetroxide containing aqueous liquid is } "neutralized" in vegetable oil, for those involved in precious metals } recycling, this act essentially kills the economics of recycling. } } 3] We have in beta testing stage right now an "osmium recycling kit". We } are looking for a limited number of laboratories (for now, just in the USA) } to participate in our beta testing of this kit. If you are interested in } participating in this test, let me know off-line and I will send you the } details. } } 4] In the mean time, I would offer the following advice. Consider using } one of the other methods described on this listserver in the past, such as } the ones involving KOH as a reducing agent. The reduced material in this } state can be recycled economically, but only in large quantities. No one } laboratory, in our opinion, could generate enough such material over a } reasonable period of time, to make recycling and refining, even of the } material is in this state, economical. } } 5] The one thing you don't ever want to do, at least in the USA, is to } declare this as any kind of a "hazardous waste". Once something has been } declared to be a hazardous waste, that designation can never (if my } understanding of regulations is correct) be reversed, and it forever has to } be treated as a waste, and translated, that means it is destined for } eventual disposal by either landfill or incineration. I want to be very } careful here, it is complicated, this is true so long as we are talking } about a RCRA hazardous waste, that is, it meets a listing or characteristic } definition. One environmental experts tells me the following: "The reason } OsO4 (and OsO2, for that matter) are not regulated as hazardous has nothing } to do with their human toxicity (or lack of); it is because osmium, from a } regulatory standpoint, is not considered toxic to the environment." He } also says it is OK to designate something as a hazardous waste with the } intent to recycle it; it simply must be managed as a hazardous waste while } it is on-site. } } So these containers that are holding reduced material (from the tetroxide) } for recycling must be labeled properly, and that would mean something like } "osmium dioxide for recycling". I am getting into an area that is not black } and white defined, and probably varies from institution to institution in } the US, not to mention the variation from country to country. But the } important thing is that from a regulatory point of view, what that label } says ends up determining the possibilities for the ultimate fate of its } contents. } } We are striving to find a way to help people transfer, on their } environmental "accounting sheets", a material from the column saying } "incineration" or "landfill", to the "recylcing" column. And for those who } worry about such things, from a legal liability standpoint, there is general } recognition that if the material is recycled, there is far less legal } liability associated with its disposal than if it is incinerated or sent to } landfill. And at the same time, recycling keeps this most valuable } nonrenewable resource in the stream of commerce and available for future } generations. } } Chuck }
There is a proposal for TEM characterisation of cadmium telluride nanoparticles for morphology and size information. I would appreciate information from the list members whether
1. CdTe and related compounds are stable under electron irradiation 2. there is a accelerating voltage limit to be adhered to 3. any other precautions required to ensure microscope safety
My primary microscopy experience is with metallic alloys and ceramics. I have this impression that these compounds are low melting, unstable and likely to sputter onto the pole pieces. Kindly correct and advise me. I use a JEOL 2000 EX II top entry stage operating at 200 kV.
---- Divakar R Physical Metallurgy Section, Indira Gandhi Centre for Atomic Research Kalpakkam 603102, India ----
Years ago, I did CdTe/CdS junction. As far as I can remember, both layers were beam sensitive and the layer CdS was more so. I could get fairly good twined CdTe structures as well as its overall morphology, but had difficulty revealing the details in CdS as the available effective observation time was much limited plus the CdS layer was very thin. There are a few ways to get around of the issue, including using a) C-coating, b) small spot size (} 2), and perhaps c) proper keV's (I was using 100keV, the max available voltage for me that time). Good luck!
Chao-Ying Ni Scientist Rodel Inc. USA
-----Original Message----- } From: Divakar R [mailto:divakar-at-igcar.ernet.in] Sent: Monday, April 09, 2001 3:58 AM To: Microscopy (E-mail)
There is a proposal for TEM characterisation of cadmium telluride nanoparticles for morphology and size information. I would appreciate information from the list members whether
1. CdTe and related compounds are stable under electron irradiation 2. there is a accelerating voltage limit to be adhered to 3. any other precautions required to ensure microscope safety
My primary microscopy experience is with metallic alloys and ceramics. I have this impression that these compounds are low melting, unstable and likely to sputter onto the pole pieces. Kindly correct and advise me. I use a JEOL 2000 EX II top entry stage operating at 200 kV.
---- Divakar R Physical Metallurgy Section, Indira Gandhi Centre for Atomic Research Kalpakkam 603102, India ----
The tripod polisher may just work. Although your sample is larger than anything I have previously tripoded, it should work if you use a plain L-bracket. First, planarize your sample and the two back feet of the tripod polisher. Then adjust the two back feet to give you the wedge angle you desire (just slightly over 1 degree if my trigonometry for your sample is correct). Likewise, the Multiprep system from Allied High Tech should be able to accept and wedge polish a sample of this size.
If this approach does not work then you may need to use a precision lapping and polishing jig. You can either design your own or you may want to check with South Bay Technology. They make several jigs with precise angular control for polishing crystals. I don’t know however, if they will work with samples this large.
Hope this helps.
Eric Windsor
Disclaimer: I have no financial interest in either South Bay Technology or Allied High Tech Products. I am a satisfied customer of both. Also, there may be other products on the market that will work equally well for preparing this sample.
The opinion expressed is my own and not that of my employer (NIST).
Original Message:
We have a professor here who has a 1 cm x 2.5cm steel layer about 100 um in thickness on a ceramic substrate. The metal layer was sputter deposited onto the ceramic substrate. The ceramic substrate extends past the metal layer. He needs to get a thickness gradient across the steel layer along the 2.5 cm length. He would like to have one end at about 50 microns in thickness and the other at 2 microns with a gradually decreasing thickness gradient. Steps down would be ok although a smooth transition would be better. We have a laser profilometer to measure anything that we produce. Does anyone out there have any ideas how to do this? Would a tripod polisher work? I thought about electropolishing and masking off portions at a time but I worry about what will happen at the interface between the ceramic and the metal. Could we alter a dimpler?
i am interested in a rapid i.e. same day processing schedule, for diagnostic } renal bx. it must be reliable and the cutting qualities good, since we do a } lot of low mag work (250x). thanks in advance. } john
{HTML} {FONT FACE=arial,helvetica} {FONT SIZE=2} i am interested in a rapid i.e. same day processing schedule, for diagnostic {BR} > renal bx. it must be reliable and the cutting qualities good, since we do a {BR} > lot of low mag work (250x). thanks in advance. {BR} > john {BR} {/FONT} {/HTML}
} In respose to Tina's post, I have not seen any } mention on the list of the scanner I purchased } a few weeks ago, the Epson Expression 1640XL. } It has 1600dpi optical resolution (scans at } a hardware resolution of 1600x3200 dpi) 42 bit } color (14 bit gray) and Dmax of 3.6.
I would certainly believe the resolution and the color depth for this scanner is adequate, but if scanning TEM films is an issue, I'd seriously advise measuring the optical density of your films ... I've heard these approach OD} 4 ... which would imply you might consider the dedicated film scanners, e.g., Polaroid 45 Ultra or the new Nikon LS-8000.
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 have been listening to the thread on scanners. Has anyone done tests of how accurate they are in absolute terms for quantitative digitization?
------------------------------------------------------- Laurence Marks Department of Materials Science and Engineering & Center for Transportation Nanotechnology Northwestern University Tel: (847) 491-3996 Fax: (847) 491-7820 mailto:ldm-at-risc4.numis.nwu.edu http://www.numis.nwu.edu http://www.ctn.northwestern.edu ------------------------------------------------------- The Other Nanotubes http://focus.aps.org/open/st12.html Boron Nitride Nanotubes http://pubs.acs.org/cen/topstory/7912/7912notw1.html
Workshop May 17-19 2001 "New approaches to the Phase Problem" http://xraysweb.lbl.gov/esg/phasing/index.html
Hello, I'm looking for companies (other than Gatan whom I've already contacted) that sell cryo-stage and cryo-prep. systems for a JEOL 5800LV scanning electron microscope. Thanks for any input! Tracey
Tracey Pepper Supervisor Bessey Microscopy Facility Iowa State University ph: 515-294-3872 fax: 515.294.1337
Has anyone used any of the denatured ethanols as a substitute for absolute ethanol?
We have recently run into some difficulty when needing to reorder 200 proof ethanol, which we use for dehydration and infiltration of samples (primarily many types of paper) prior to embedding in Spurrs epoxy, and for cleaning samples (non paper) and lenses of light microscopes. The chemical company selling the ethanol is insisting that we must have a liquor license before they will ship to us.
Ethanol denatured with a variety of substances is readily available and can be shipped with no licensing requirements. Our concern is that the denaturing agent will leave a detectable residue on lenses, samples, and may cause problems with the polymerization of Spurrs. Rather than obtaining a liquor license, we are considering using one of the 100:5 ethanol: methanol blends. If any of you have had successful or unsuccessful experiences substituting denatured ethanol for absolute in embedding or cleaning protocols, I would appreciate hearing from you.
Teresa Boes Hewlett-Packard Analytical and Development Lab 1000 Circle Blvd Corvallis, OR 97330 541-715-7055 teresa_boes-at-hp.com
Sorry about being late with this thread...Out here we do about 750 renal biopsies last year and all our sections are mounted on 300 mesh uncoated thin bar Gilder grids. We have a Philips 208S at 80kV. We can get a majority of the glomerulus to lie in the grid square to be viewed. Most of our images are shout between 2800X and 14,000X.
Last November we finally obtained the AMT Advantage HR 1K x 1K system and use it exclusively for our EM images... We also do some Neuropathology and Surgical Pathology EM in this lab.
Out here we have rigged up a system that all the Pathologists who need the EM images are setup on a EM users group on the network. I them upload the images from the computer here in the scope room to a directory on our network so the Pathologists can view the images right at their desktop. The renal Pathologist here is thrilled with the system... It cuts down our expenses, and it shortens our turnaround time on specimens from 5 days to about 3 days or less....
Eric A. Rosen Electron Microscopist UCLA Medical Center
==============================
} John: } We do around 500 renal biopsies per year and all the sections are } mounted on 200 mesh uncoated copper grids. We have an 8 year old Hitachi } 7100 and use 60kv. The majority of the glomerulus can be viewed with the } 3-4 serial sections lying randomly across the grid bars. We do not need a } picture of the whole glomerulus, rather most pictures are between 3,000 and } 10,000X. } Dr. Tibor Nadasdy is the renal pathologist and decided last year that } all our renal biopsies would be captured with the digital camera onto a } computer and sent up to him via a network to his computer. So, at the } present time we use very little EM film. He diagnoses each biopsy and } e-mails representative digitized images to the nephrologists. } } } } } Karen L. Jensen, M.S. } Project Manager & Associate Scientist } Electron Microscopy Research Core } -----Original Message----- } } From: "JHoffpa464-at-aol.com"-at-sparc5.microscopy.com } [mailto:"JHoffpa464-at-aol.com"-at-sparc5.microscopy.com] } Sent: Friday, March 30, 2001 2:20 PM } To: microscopy-at-sparc5.microscopy.com } Subject: renal Em } ok taking a little survey. i am in a diagnostic EM lab. we mount out } sections } on formvar coated slotted grids, so he can shoot pics of the whole } glomerlus. } ok my question. how may of you out there doing diagnostis EM on renals do } this? } john
Tracey VG Microtech make Polaron integrated column-mounted cryo systems including a new system designed for high-resolution work with FEGSEMs Emitech have an off-microscope cryo-prep and transfer system suitable for LTSEM with a conventional tungsten filament or LaB6 SEM BalTec make various cryo transfer and preparation systems which could be used to transfer specimens from e.g. a freeze-etcher to SEM
see http://www.kaker.com/mvd/list.html for addresses and contact numbers for these companies Chris
----- Original Message ----- } From: "Tracey M. Pepper" {tpepper-at-iastate.edu} To: {microscopy-at-sparc5.microscopy.com} Sent: Monday, April 09, 2001 6:52 PM
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I am looking for a 2" and/or 3" scribe tool for GaN. Does anyone have any experience with a quality new/used equipment vendor who supplied such a tool in the past. I am looking for a fairly new programmable scriber.
Eric, I think that your geometry is off. The Tripod Polisher is about 50 mm long from the sample to the feet and has 1 um divisions. The arctan of 1/50000 gives about 0.001 degrees. This job needs an angle given by the arctan of 48/25000 or 0.11 degrees. Plus you need to stick the thickness at one end at a particular thickness -2 um. It is doable with the TP, but will be difficult. You are correct that they will need to planarize the sample. What I would do is also planarize the holder and mount the parallel-sided sample and then set the height of the feet to the thickness of the sample and then add the amount for the angle. Then I would slowly polish using a low value grit 1 or 3 (perhaps even 1/2) um until I went through to the substrate at one end. You would have your angle and thickness values that went from zero to the desired value and a little thicker. The trick is stopping at the right place and accounting for the wear on the feet. You should be able to watch the facet move towards one end. You can watch the progress using a glass plate and look at the thickness fringes at the facet caused by the unpolished and polished surfaces.
-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)
"The opinions expressed are those of Scott D. Walck and not of PPG Industries, Inc. nor of any PPG-associated companies." --
} -----Original Message----- } From: Eric Windsor [mailto:Eric.Windsor-at-nist.gov] } Sent: Monday, April 09, 2001 9:15 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: Re: Preparation of a steel wedge on a ceramic substrate } } } -------------------------------------------------------------- } ---------- } 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 } } -------------------------------------------------------------- } ---------. } } } Robin, } } The tripod polisher may just work. Although your sample is } larger than } anything I have previously tripoded, it should work if you use a plain } L-bracket. First, planarize your sample and the two back feet of the } tripod polisher. Then adjust the two back feet to give you } the wedge angle } you desire (just slightly over 1 degree if my trigonometry } for your sample } is correct). Likewise, the Multiprep system from Allied High } Tech should } be able to accept and wedge polish a sample of this size. } } If this approach does not work then you may need to use a } precision lapping } and polishing jig. You can either design your own or you may } want to check } with South Bay Technology. They make several jigs with } precise angular } control for polishing crystals. I don't know however, if they } will work } with samples this large. } } Hope this helps. } } Eric Windsor } } Disclaimer: I have no financial interest in either South Bay } Technology or } Allied High Tech Products. I am a satisfied customer of } both. Also, there } may be other products on the market that will work equally well for } preparing this sample. } } The opinion expressed is my own and not that of my employer (NIST). } } Original Message: } } We have a professor here who has a 1 cm x 2.5cm steel layer } about 100 um in } thickness on a ceramic substrate. The metal layer was sputter } deposited } onto the ceramic substrate. The ceramic substrate extends } past the metal } layer. He needs to get a thickness gradient across the steel } layer along } the 2.5 cm length. He would like to have one end at about 50 } microns in } thickness and the other at 2 microns with a gradually } decreasing thickness } gradient. Steps down would be ok although a smooth transition } would be } better. We have a laser profilometer to measure anything that } we produce. } Does anyone out there have any ideas how to do this? Would a tripod } polisher work? I thought about electropolishing and masking } off portions at } a time but I worry about what will happen at the interface } between the } ceramic and the metal. Could we alter a dimpler? } } } Thanks, } Robin Griffin } UAB } } }
How you can get the licence if you do not SELL the alcohol? Licence is only for selling..
} We have recently run into some difficulty when needing to reorder 200 proof } ethanol, which we use for dehydration and infiltration of samples (primarily } many types of paper) prior to embedding in Spurrs epoxy, and for cleaning } samples (non paper) and lenses of light microscopes. The chemical company } selling the ethanol is insisting that we must have a liquor license before } they will ship to us.
Does anybody have experience (good or bad) with Imaging Plate scanners on TEMs?
PLease reply on or off-line, I will post a summary - preserving anonymity if necessary!
thanks
Sally
Dr Sally Stowe Facility Coordinator, ANU Electron Microscopy Unit Research School of Biological Sciences Australian National University Canberra ACT0200 AUSTRALIA stowe-at-rsbs.anu.edu.au fax 61 (0)2 6125 3218 or 6125 8525 http://www.anu.edu.au/EMU
Your point is well taken, but in this environment, there is no distinction between purchase, application or consumption.
I'm in California and have not run into this situation as yet. But I would bet that the same myopic rules would apply. I buy ethyl alcohol from Ted Pella in Redding, CA. But I don't know the specific proof of the alcohol (Cat.# 19207). It comes in 200mL bottles and poses no problem when purchased in lots of six or fewer bottles.
gg
At 03:38 PM 4/9/2001, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
How do you define "quantitative digitization?" i.e., what variables are you dealing with in this respect? What are the "absolute terms?"
Anyone else have some ideas about this topic?
gg
At 10:10 AM 4/9/2001, you wrote:
} I have been listening to the thread on scanners. Has anyone done } tests of how accurate they are in absolute terms for quantitative } digitization? } } ------------------------------------------------------- } Laurence Marks } Department of Materials Science and Engineering & } Center for Transportation Nanotechnology } Northwestern University } Tel: (847) 491-3996 Fax: (847) 491-7820 } mailto:ldm-at-risc4.numis.nwu.edu } http://www.numis.nwu.edu http://www.ctn.northwestern.edu
I am a Laboratory Technician currently studying for a Diploma in Occupational Health and Safety. My last study module is an Action Research project which I would like to do on the development of ergonomic microscopes and although there is a lot of information on the problems involved in microscopy and methods to relieve them I was wondering if any research has been done into the effectiveness of ergonomic design for improving microscopy diagnosis, and where I could get any papers on the subject. Thanks. Andy.
Perhaps the reason there was no recipe for the GMA is that you can vary the component ratios alot depending on the tissue you're embedding. There are several reviews from the 60s and 70s on different GMA recipes, at least for embedding plant material. One "standard" mix for plant material is 95% v/v pure GMA, 5% v/v polyethylene glycol, 0.15-1.0% w/v benzoyl peroxide (catalyst). With more catalyst, polymerisation is faster and blocks are harder, but too much produces brittle, bubbly blocks. PEG can be 0-10%, PEG 200 and 400 are commonly used.
Like other methacrylate resins, GMA can be heat polymerised but you need to exclude oxygen. You can either seal the GMA blocks in capsules - gelatin capsules for example - dent the cap to exclude as much air as possible, or cover flat embedding moulds so that they are completely sealed - e.g. use a plastic vial cap as flat mould with another on top to seal. If you have access to a vacuum oven, you can polymerise open moulds under nitrogen.
Time to polymerise depends enormously on the composition of the resin, try 60C overnight for starters if using a fairly "standard" mixture.
good luck, cheers, Rosemary
Rosemary White Microscopy Centre CSIRO Plant Industry GPO Box 1600 Canberra, ACT 2601 Australia
Just a minor correction / update. VG Microtech recently changed name to Thermo VG Scientific, address and contact details remain the same, except for the surface science website:
The Polaron range continues to be represented in the US by Energy Beam Sciences.
Best regards,
Mike Wombwell Polaron Range Business Manager Thermo VG Scientific The Birches Industrial Estate Imberhorne Lane East Grinstead West sussex RH19 1UB UK Tel: +44(0)1342310296 (direct line) Tel: +44(0)1342327211 (Switchboard) Fax: +44(0)1342315074 email: mike.wombwell-at-scientific.com Website: http://www.polaron-range.com
E & OE
-----Original Message----- } From: Chris Jeffree [mailto:c.jeffree-at-ed.ac.uk] Sent: 09 April 2001 22:04 To: Tracey M. Pepper Cc: microscopy-at-sparc5.microscopy.com
Tracey VG Microtech make Polaron integrated column-mounted cryo systems including a new system designed for high-resolution work with FEGSEMs Emitech have an off-microscope cryo-prep and transfer system suitable for LTSEM with a conventional tungsten filament or LaB6 SEM BalTec make various cryo transfer and preparation systems which could be used to transfer specimens from e.g. a freeze-etcher to SEM
see http://www.kaker.com/mvd/list.html for addresses and contact numbers for these companies Chris
----- Original Message ----- } From: "Tracey M. Pepper" {tpepper-at-iastate.edu} To: {microscopy-at-sparc5.microscopy.com} Sent: Monday, April 09, 2001 6:52 PM
Richard: Not true in the good old US of A! 200 proof EtOH is considered a controlled substance, and purchase of even small quantities is regulated. In the Histology and EM Labs here, we use a fairly large supply, so there is a company license which allows purchase of a specific amount, and that means if demands go up dramatically there is a real issue of obtaining adequate supplies from time to time. I think that the issue may well have to do with the quantities being purchased. If one buys the odd bottle now and then, there is no hassle. But when usage passes a certain point, then licenses are necessary. Keeps all of the bureaucrats in jobs. As for the license, Teresa, it is mostly just paperwork. Requires you to specify purpose for use, amounts that will be required, etc. If I remember correctly (had to do this myself a _very_ long time ago--now an internal supply room clerk does it), this is an annual process. And, like most labs, since you have to account for every drop of reagent that enters and leaves the lab for EPA, state EPA, OSHA, etc, you will likewise enter the EtOH into that stream, thus showing that you aren't using it for more pleasurable ends.
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals NB--personal opinions and experiences only expressed above. On Tue, 10 Apr 2001 08:38:10 +1000, Vr. Richard Bejsak-Colloredo-Mansfeld 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 | -----------------------------------------------------------------------. | | | How you can get the licence if you do not SELL the alcohol? | Licence is only for selling.. | | } We have recently run into some difficulty when needing to reorder 200 | proof | } ethanol, which we use for dehydration and infiltration of samples | (primarily | } many types of paper) prior to embedding in Spurrs epoxy, and for cleaning | } samples (non paper) and lenses of light microscopes. The chemical company | } selling the ethanol is insisting that we must have a liquor license before | } they will ship to us. | | |
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc. 900 Rigdebury Road Ridgefield, CT 06877 203-798-5448
_______________________________________________________ Send a cool gift with your E-Card http://www.bluemountain.com/giftcenter/
I am interested in freeze drying pieces of murine liver tissue for subsequent embedment in Spurr's epoxy resin. Does anyone have experience with this technique? Specifically I would be interested in times and temperatures. I will be using a turbo freeze dryer from EMS to freeze dry the tissue.
Thanks in advance,
Germaine G. Boucher TEM Lab Pfizer Global Research and Development Groton, CT
Would noise be a good criterion? Say, for a perfectly evenly darkened film (if such a thing existed, or at least even on a scale { { collected pixel size) - what is the value of the noise (standard deviation of pixel value) as a function of film darkness (density)?
This would presumably improve with the time of collection. Thus how "good" your scanner is depends on how you run it or whether it lets you take a slower scan or to average multiple scans. With the exception of drum scanners these devices all use CCD arrays. So what is probably most of interest is the signal to noise ratio as a function of illumination intensity, with everything known about CCD's going into determining this. The maximum density the scanner can handle is just the point at which the noise swamps the signal.
There must be some good literature out there on the sources of noise, optimizing collection (scan) time etc. One article which might be a starting point is:
G. H. Campbell, W. E. King and D. Cohen "Analysis of Experimental Error in High Resolution Electron Micrographs", Microscopy and Microanalysis vol. 3 (1997) p. 451.
This is not very detailed, and treats only the total random noise, i.e. grouping noise arising in collecting the image with that arising from the scanner.
Now, finding a good "Consumer Report" test with hard numbers on commercial models is likely to be a lot harder!
Wharton
} -----Original Message----- } From: Gary Gaugler [SMTP:gary-at-gaugler.com] } Sent: Monday, April 09, 2001 10:37 PM } To: L. D. Marks } Cc: MSA listserver } Subject: Re: Scanners: quantitative accuracy } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } How do you define "quantitative digitization?" i.e., what } variables are you dealing with in this respect? What are } the "absolute terms?" } } Anyone else have some ideas about this topic? } } gg } } } At 10:10 AM 4/9/2001, you wrote: } } } I have been listening to the thread on scanners. Has anyone done } } tests of how accurate they are in absolute terms for quantitative } } digitization? } } } } ------------------------------------------------------- } } Laurence Marks } } Department of Materials Science and Engineering & } } Center for Transportation Nanotechnology } } Northwestern University } } Tel: (847) 491-3996 Fax: (847) 491-7820 } } mailto:ldm-at-risc4.numis.nwu.edu } } http://www.numis.nwu.edu http://www.ctn.northwestern.edu }
To the person having trouble getting 200 proof ethanol:
If you are embedding in Spurrs, you do not need ethanol or P.O.. Just dehydrate in E M grade acetone and infiltrate with acetone/Spurrs and you will get good results. Keep it simple, Tim
Timothy G. Schneider Director of Electron Microscopy Department of Pathology Room 229 Jefferson Hall Thomas Jefferson University 1020 Locust St. Philadelphia Pa. 19107 215-503-4798 work 610-613-8170 cellular timothy.schneider-at-mail.tju.edu
I have put together a summary of responses I got from my inquiry about a week or so ago about EDS systems. I really appreciate everyone's input. We haven't made any decisions yet since we are still gathering information but your responses will help. I've included the responses in their entirety so I hope this helps others as well.
Thanks again from everyone who contributed.
Jean Ross Central Microscopy Research Facility University of Iowa
----------------------------------------------------------------------------- I have been using an IXRF EDS / Gresham Detector (with IXRF digital pulse processor) system for almost 2 years. Initially, the IXRF was installed on an ETEC. About 15 months ago the Etec was replaced with a Hitachi 3500, a new detector (Gresham) was purchased, and the EDS was installed on the 3500. Generally, I am quite satisfied. There are minor software bugs, but IXRF has been reasonably good at fixing them when discovered. It has been my experience that all the systems have bugs, perhaps some more than others. Prior to the IXRF, I had a Kevex 8000/Delta.
Low end noise and broad peaks were evident on first installation, but were soon fixed by tweaking the detector preamp and pulse processor amp.
I am still running their first software package, "Iridium". I have the newest release, "EDS 2000", but lack of time has kept me from installing and checking it out.
I should mention that IXRF is a "virtual" company, with people spread out between Texas, California, etc. This has not proven to be a problem.
Woody White McDermott Technology, Inc. nwwhite-at-mcdermott.com
We have had EDAX for about a decade and a half, and we are very pleased with the product and the service we get. Carol Heckman heckman-at-bgnet.bgsu.edu
I would recommend you give consideration to Doug Connors at TN Analyzer Service, Inc. of Dane, WI. Doug has rebuild and upgraded detectors for me for the last 6 years. He is dependable knowledgeable, and economical as well.
Bob Roberts EM Lab Services, Inc. 2409 S. Rural Rd Suite C Tempe, Arizona 85282 (480) 967-3946 bobrobs-at-earthlink.net
We recently purchased a Noran Instruments Vantage DS1. We purchased this based on some very impressive demonstrations of software that the salesperson brought in. Unfortunately they are still working out the bugs in their software. Everything they have is ported over from Unix, and literally runs in a unix shell on an Microsoft Windows NT platform. This makes their software fairly buggy. Their response time to fix major bugs and hang-ups in the software has been very slow, and if given the opportunity to do it all over again I'd probably look at Oxford Instruments. I would still rate the quality of the equipment very high. Our detector performs at the specified resolution, and is a good piece of equipment. Now if they could only get the software end of it straight... My vote: 1) Oxford Instruments 2) Noran Instruments 3) Edax or some of the smaller players The benefit with going with a larger company is support and upgrades. We have a 10 year old WDX that we just purchased new software and interface for last year. Our old EDS was given some trade in value by Noran. And we all know how valuable a service contract can be...
Get back to me if you have any more questions, ~Jonathan Jonathan Dunlap Analytical Laboratory Manager Osram Sylvania Inc. 816 Lexington Avenue Warren, PA 16365 Ph: 814-726-6991 Fax: 814-726-6942 Jonathan.Dunlap-at-sylvania.com
We have an Oxford Instruments Link ISIS Model 200 on our 2460N. We have been happy with it, but I don't know about the direction that Oxford is heading. I don't care for the feel of their new INCA software. Some might like it. It also seems to be slow coming together. Some of the functions are still lacking after 2 (or is it 3) years of seeing it at MSA.
We have an IXRF system on our JEOL 840A. It was a good price ($30K) for an upgrade to our Kevex several years ago. It does what we need. They keep at work on the software and have it freely available on the web. I might have to pay closer attention and stay away from the beta stuff. They are still working on it. They also have a nice digital pulse processor which stills stand alone for about $5k.
I still feel funny about some contacts with EVEX. I can't say much about EDAX, NORAN, or PGT. They should all have good stuff but it might be pricey. The last we seriously looked at them was 6 years ago or so when we opted for the Oxford.
I was intrigued by the unit from Quartz PCI. I think it was called X-ray One, or such. It was new at MSA 1-1/2 years ago but looked promising.
We purchased an EDAX Falcon system for our Hitachi S-3000N and I've been pleased with it. It has better light element sensitivity than most which was very important to me although I don't think that its mapping capabilities are as good as PGT's, say. I don't have direct experience with Noran although I did talk to them and their system seemed ok - but logistics didn't favor Noran so I passed on them. EDAX does have good integration with the Hitachi and the Quartz database.
I'd be glad to respond more specifically if you'd like.
Richard Shalvoy Arch Chemicals Cheshire, CT RBShalvoy-at-archchemicals.com
I have an iXRF systems out of Texas using a Gresham detector. It works well. Not the most cutting edge, but they are one of the "start ups". They have been around for I guess 6-7 years. I have a digital pulse processor and completely active control for x-ray maps and such. They are very price competitive, but lack a dedicated technical support person. You talk to the programmer or electronics expert, but no techs on the phone whenever you have a software question. But, if you willing to wait a day for some answers then they are worth it. I haven't run across the problem where I thought, "if I just had a better system". If you want to integrated w/ WDS than maybe Noran. Also, if you want to integrated w/ motorized stage control, I don't think they off such a package, like the bigger companies.
I have a Hitachi 450. I used to run a 2400 and 500 before I quit my day job and went out on my own. I am very happy w/ Hitachi.
I use Oxford ISIS300 system on HITACHI S-3500N (with VP mode) for light element analysis, mostly C, O, N, F, P, S, Si, Mg, as well as metal Co, Ni-P, Pt, Cr, Fe, W, etc. This system works well. One useful function is the overlay of 2 spectrums. I can easily subtract the blank from the sample spot and make it easy to identify what is (are) in the sample. I am sure some other program may have this kind of function, but I have not seen.
Zhiyu Wang zhiyuw-at-home.com I would be interested in seeing the responses as I am going to try and get funding next year for a replacement for our EDAX PV9100 on an Hitachi s-450.
We have been running EDX on SEMs and TEMs for many years. We used to have a range of systems from Kevex, PGT, Noran, Link, EDAX, however a few years ago we decided that we ought to standardize on one common system. After evaluation we bought three Oxford Instruments ISIS systems. Whenever we have upgraded or bought new systems they have been Oxford Instruments ISIS or now INCA.
I have been happy with the ISIS except for the file handling that was not designed for a multi user facility such as ours (approx 120 EM users in total roughly 25 to 30 swapping every year). I am really quite impressed by the INCA, Oxford Instruments are, at last, listening to the users and adding user requested facilities. They have sorted out the file handling mess of the ISIS and structured it well for an SEM user (not quite as well for a TEM user but there are less of us). The software structure is quite intuitive and there is a really impressive help menu and explanation of everything from the physics of X-ray generation, how EM’s work, how detectors work and how to analyze samples.
Their detectors have always been good and the SATW (thin window detectors) still have a reasonable efficiency at low Z. B is possible but C is easy and even the N peak is over 30% efficient (there is often a high absorption at N).
Another feature that is invaluable for TEM is the integral shutter that will close when the count rate is too high. This protects the crystal, it prevents it overloading and shutting down or worse the crystal efficiency may change for a few minutes until it recovers fully. This may affects your quantitative work. In TEM this is usually caused by hitting the grid bar and not really a problem in SEM but I don't know what secondaries and ions you will have in a variable pressure SEM. It could be useful for you, check with other high pressure SEM users.
Regards, Ron
Please note: Oxford Instruments have upgraded an ISIS to an INCA system in my department, without charge, in return for access to the instrument for development projects and demonstrations for a fixed number of days. I receive no benefit from this and the department has no benefit from Oxford Instruments sales. I remain a thorn in the flesh of all our suppliers if I think they could improve their products or service. ron.doole-at-materials.oxford.ac.uk
Hello, I am very familiar with the Oxford ISIS 300 series spectrometers. They are ok, and the new Inca system looks good too. However, I recently saw the PGT spectrometer at Lehigh and it is very impressive.
I'm also in the market for an EDS system and have looked at EDAX, Noran, PGT and Oxford.
I edited out PGT because in order to quantify you have to optimize the system for the type of sample by playing with fudge factors, which none of the other systems have to do (though one of them, I think Noran, lets you adjust a sensitivity factor if you want to, but they didn't do it on my samples that were tested against known microprobe results and the answers were fine). I also eliminated Oxford, though it has a terrific user interface (maybe at the expense of functionality), because they consistently IDed my aluminum peaks as Br or Tm (!); this made me wonder about all their algorithms. They claim it had to do with the takeoff angle on the particular SEM being used, but that shouldn't be a factor.
I like both EDAX and Noran, though for different reasons. EDAX user interface is better than Noran's, though again, I think Noran possibly offers more routines (it's hard to keep track and see absolutely everything a system has to offer in a demo day....).Noran can multitask - work on several programs while a spectral map is being collected, for instance (does EDAX? I have to check). But EDAX has a beam skirt reduction routine for low vac mode (though it's time consuming, so a bit cumbersome), and their peak modeling is right up front - but Noran can put theirs up front also if you want to have it accessible (yes) and I think Noran might be a little better engineered.
As you can see I'm still in a quandary (ditto for the two contending SEMs, LEO and ESEM). Whatever I decide I'll still be very interested in the results of your posting - especially if other folks' info comes in within the next week or so it would help in my decision too.
I hope my input helps a little. Good luck with your quest!
I looked into Noran, EDAX, and PGT. Noran was quickly culled (less user friendly, less abilities, didn't work right during demo), but EDAX and PGT both seemed to have equivalent capabilities (PGT claimed a 'proprietary' signal amplifier/digitizer doohickey, but it was only in the placement.) For the long-time spectrum gathering (I forget the technical term), PGT makes many passes with short dwell times while EDAX dwells on each pixel much longer to collect data & does it in 1 pass. Kinda 6 of one, half dozen of the other. What made us choose the EDAX Phoenix system was the fact that the PGT software was UNIX-based (although hidden) while EDAX is PC-based. I've heard rumors that PGT is switching to PC-based; we purchased our system in 1999. I've also heard that Noran has practically no service techs (but that may be an East coast thing.) We've been happy with the EDAX service, and I enjoyed their user school very much. By the way, our SEM is a variable pressure JEOL 5900, and it's integrated with the EDAX system.
Hope I've been helpful,
Jane L. LaGoy Development Engineer Bodycote IMT, Inc. 155 River Street Andover, MA 01810 978-470-1620 jlagoy-at-bodycote-imt.com
Hi All: I have been asked to examine some films on a Si carbide substrate using TEM. Does anyone out there work with this material. Can one mechanically thin it using diamond films? Please offer some suggestions of what you are doing. Thanks in advance, Michael Coviello University of Texas Arlington
We have some staining that suggests the staining dye has transferred from one place to another and I write to see if anyone could shed some light on this for us.
In the experiment we stain isolated chromosomes with DAPI, rinse them, and then introduce them into plant protoplast cells (likely one or few per cell). There is no autofluroescence in the DAPI channel and we can follow the course of the dyed chromosomes over time. At first the DAPI fluorescence appears in the cytosol on structures likely to be the introduced chromosomes. Then it appears in the nucleus, where all the chromosomes are stained! This is especially evident when we culture these cells. Dividing cells at metaphase show DAPI fluorescence over the entire metaphase plate. Note that only some cells show DAPI fluorescence, consistent with the presumption that our fusion process that introduces the chromosomes is only successful in some of the cells.
Has anyone had an experience where dye has been transferred from one structure to another in a living cell? What are some alternative interpretations of this phenomenon?
Thanks for your comments,
Howard Berg
R. Howard Berg, Ph.D. Director, Integrated Microscopy Facility, Associate Member Donald Danforth Plant Science Center/Nidus Center 893 North Warson St. Louis, MO 63141
We tripod polished thin films on SiC. It works OK with diamond lapping film and diamond powder slurry as a final polish, but it takes a loooonnng time. Put a sacrificial piece of SiC on the second side tool pedestal so the final wedge polish will go from SiC sample to SiC tool. Otherwise the glass pedestal polishes faster than the SiC and makes a little step that can lead to sample breakage. ...Always a good idea to match the pedestal hardness to the specimen--hard or soft.
As I recall, we had more problems with poor film adhesion to the SiC than with polishing the SiC substrate.
Although we haven't tried it, I suppose a FIB would work fine.
Ron
There was a thread recently on scanners for TEM film. I have looked up all the models mentioned, on the web and called agents for prices - and produced a comparative table, given below.
I do not guarantee that the figures are accurate but they are my best interpretation of the data given.
In the light of experience and Nestor's comments, I would suggest that 2000 dpi is a minimum for TEM negatives. You may be able to get away with less nine times out of ten, but there will be occasions when you need more. I would exclude the Minolta and all the Epsons from consideration (despite the incredibly low prices of some of the Epsons) because of the low pixel density.
Among the rest the Nikon has the best pixel density and the best optical density (another critical parameter for TEM negatives). The price is very competitive too. The Nikon web site does not give a time for scanning a negative. On the face of it the Nikon would be a best buy - get a separate, inexpensive flatbed scanner for the other work.
These comments are all my own opinions based on manufacturers' data. Since we are considering purchase any comments to the contrary would be most welcome.
Code Maker Model Type
A Agfa DuoScan T2500 Flatbed -Transparency included
B Epson 1640 several versions Flatbed -Transparency option 1680 several versions
C 1600 several versions Flatbed -Transparency included
D Imacon Flextight Precision II Drum -for film and large format
E Minolta Dimage ScanMulti II Film
F Nikon Super Coolscan 8000ED Film
G Polaroid 45 Ultra Film
H Umax Powerlook 3000 Flatbed -Transparency included
Code dpi OD Time Price Opinion at 6 x 9 cm
A 2500 x2500 3.4 3 min $4,500 Fair
B 1600 x 3200 3.6 $300-$3000 Poor $800-$1400 Poor
C 1600 x 3200 3.3 $650-$1160 Not suitable
D 2240 x2240** 3.9/4.1 N/A above $10k Good: low pixel density
E 1128 x 1128 3.6 Not suitable
F 4000 x 4000 4.2 N/A $2,695 V. Good
G 2500 x 2500 3.8 5 min $7,495 Good but pricey
H 3048 x 3048 3.6 3 min $6,499 Good
-- .......... Alwyn Eades Department of Materials Science and Engineering Lehigh University 5 East Packer Avenue Bethlehem Pennsylvania 18015-3195 Phone 610 758 4231 Fax 610 758 4244 jae5-at-lehigh.edu
I too am about to buy and I would make a couple of comments on your evaluation. First, let me remind everyone that the Dynamic range is a log scale so small numerical differences are significant.
I also think the Nikon Coolscan 8000 looks great but it only takes a 2.5 x 3.5 negative which is smaller than my JEOL and Hitachi EM negative sizes (~ 3 1/2 by 4 1/2"). Have these EM manufacturers gone to a smaller film size or is Nikon using a non-Japanese EM as their standard? seems odd but I don't see how the Nikon would be very useful. You say a {2000 line scanner would be useful 9 out of 10 times but want the 2000+ lines for the occasional high res scan. I would argue that the size of the negative was the more important variable to be worried about. The Nikon couldn't handle 4x5 LM negatives or transparencies from autoradiography of Westerns/Northerns, etc.
My leading candidate is the ArtixScan 1100 has a Dmax of 3.9 (about $1600 with SCSI card). This was has a 1000 x 2000 dpi resolution. more details at www.microtek.com. This is my leading candidate. It was 4 negative carriers and I await word whether one could be modified to carry a 3 1/2 by 4 1/2 negative. At worst, I will have my scientific instrumentation shop guys fabricate a holder. It comes with a glass 8 x 10 glass carrier for odd size negs but I want to avoid Newton rings and want a glassless carrier.
I would appreciate comments on the following argument (I think I have this correctly figured out but am not sure since so many out there seem to want to have a higher resolution scanner). I have a Fuji Pictrography 3000 printer with a 400 dpi output that is as good as any other widely available printer in the academic world. If you figure the maximum published image size is about 8 inches, that would mean the maximum image size be 3200 dpi wide. A 1000 dpi scan of my negative would be 4500 x 3500 dpi. I could crop by about 28% or 10% depending on the orientation of the negative and still be taking full advantage of the printer resolution. In reality, most EM publication prints are smaller than 8" wide so one could crop even more and still not need more than 1000 dpi. A resolution } 1000 dpi would be useful for subtle morphometric analysis but a 4000 dpi scan of a 3 x 4 negative would be 192 MB. That is pretty big for doing morphometry on! A 1000 dpi scan of a 3.5 x 4.5" negative would be about 16 MB and that is much more manageable. Perhaps the difference is in the type of EM we are doing. I am working with biological specimens doing standard thin section type stuff. are you doing some Material Sci application that demands more?
I will be interested in Alwyn (and any others) reply since I hope to buy one soon!
} . } } } There was a thread recently on scanners for TEM film. I have looked up } all the models mentioned, on the web and called agents for prices - and } produced a comparative table, given below. } } I do not guarantee that the figures are accurate but they are my best } interpretation of the data given. } } In the light of experience and Nestor's comments, I would suggest that } 2000 dpi is a minimum for TEM negatives. You may be able to get away } with less nine times out of ten, but there will be occasions when you } need more. } I would exclude the Minolta and all the Epsons from consideration } (despite the incredibly low prices of some of the Epsons) because of the } low pixel density. } } Among the rest the Nikon has the best pixel density and the best optical } density (another critical parameter for TEM negatives). The price is } very competitive too. The Nikon web site does not give a time for } scanning a negative. On the face of it the Nikon would be a best buy - } get a separate, inexpensive flatbed scanner for the other work. } } These comments are all my own opinions based on manufacturers' data. } Since we are considering purchase any comments to the contrary would be } most welcome. } } } } Code Maker Model Type } } } } A Agfa DuoScan T2500 Flatbed } -Transparency included } } B Epson 1640 several versions Flatbed } -Transparency option } 1680 several versions } } C 1600 several versions Flatbed } -Transparency included } } D Imacon Flextight Precision II Drum -for } film and large format } } E Minolta Dimage ScanMulti II Film } } F Nikon Super Coolscan 8000ED Film } } G Polaroid 45 Ultra Film } } H Umax Powerlook 3000 Flatbed } -Transparency included } } } } } } Code dpi OD Time Price } Opinion } at 6 x 9 cm } } } A 2500 x2500 3.4 3 min } $4,500 Fair } } B 1600 x 3200 3.6 } $300-$3000 Poor } } $800-$1400 Poor } } C 1600 x 3200 3.3 } $650-$1160 Not suitable } } D 2240 x2240** 3.9/4.1 N/A above } $10k Good: low pixel density } } E 1128 x 1128 3.6 } Not suitable } } F 4000 x 4000 4.2 N/A } $2,695 V. Good } } G 2500 x 2500 3.8 5 min } $7,495 Good but pricey } } H 3048 x 3048 3.6 3 min } $6,499 Good } } } -- } .......... } Alwyn Eades } Department of Materials Science and Engineering } Lehigh University } 5 East Packer Avenue } Bethlehem } Pennsylvania 18015-3195 } Phone 610 758 4231 } Fax 610 758 4244 } jae5-at-lehigh.edu
-- 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)
I assume that your SiC is single crystal. The small angle cleavage technique works for SiC. See MRS Proceedings, TEM Prep IV Vol 480.
-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: Michael Coviello [mailto:coviello-at-mae.uta.edu] } Sent: Tuesday, April 10, 2001 10:59 AM } To: listserver } Subject: TEM-SiC wafer sample prep? } } } -------------------------------------------------------------- } ---------- } 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/MicroscopyLists } erver/FAQ.html } } } } -------------------------------------------------------------- } ---------. } } } Hi All: } I have been asked to examine some films on a Si carbide } substrate using } TEM. Does anyone out there work with this material. Can one } mechanically thin it using diamond films? Please offer some } suggestions } of what you are doing. } Thanks in advance, } Michael Coviello } University of Texas Arlington } } }
Microscopy Scientist for two Recently Established NIH Centers for Biomedical Research at The University of Wyoming
As part of our seven interrelated projects into the biology, chemistry, and molecular biology of cardiovascular function and nitric oxide we seek a skilled microscopist to be hired at the non-tenured Assistant Research Professor level. Experience and research interests in state-of-the-art microscopy, including confocal and epifluorescence, and ultrastructural techniques. Primary responsibility will be management of the University's Microscopy Center. Opportunity includes the possibility of establishing a research program within this Center. Appointment will be at a (non-tenure track) Assistant Professor level for a renewable four year term.
Job Description: The person we seek will be responsible for organization of a new research laboratory facility at the University of Wyoming which will include two confocal microscopes and an electron microscope. The position includes overall management of the microscope facility, user training, and user supervision. Requirements for the position include experience with light, confocal, and transmission electron microscopy. This individual will oversee all aspects of specimen accession and processing, operation of the microscopes, photography, and record keeping. The individual will work with only minimum supervision. Responsibilities include: Serve as the technical manager of the facility and be responsible for the operation and maintenance of the confocal and EM microscope facility. In addition the manager will perform preventative maintenance on the equipment; maintain the lab, order supplies, schedule instruments, and oversee billing. Image analysis at the light, confocal and electron microscopic levels and preparation of micrographs for publication. Applicant Qualifications: Experience focus on both confocal and EM. Regarding confocal microscopy, we require experience with confocal and digital imaging techniques, visualization of living cells containing fluorescent probes, photobleaching, and fluorescence in situ hybridization. The successful applicant will have experience with tissue preparation for EM and the maintenance of an electron microscope. Excellent interpersonal and organizational skills are essential. Ph.D.. degree required Desirable Experience: Expertise and training in the operation of confocal microscope and EM microscope systems is required. Familiarity with light microscopy methods, immunofluorescent staining, use of fluorescent probes for physiologic measurements and the general principles of cell biological research are desirable. Significant facility with computers is desired. Salary Range: Commensurate with experience.
For additional information see our websites: www.uwyo.edu/nocobre www.uwyo.edu/MolecBio/Cobre To apply send complete CV, three references, and a cover letter indicating which position(s) you are applying for to: Lynda Payne, Department of Chemistry, University of Wyoming, Laramie, WY 82071-3838, USA. The searches will remain open until all positions are filled.
The University of Wyoming is located in a high (2,200 m) valley surrounded by the Rocky Mountains in the southeast corner of Wyoming. The University of Wyoming is an equal opportunity/affirmative action employer.
The Beam, newsletter for the Southeastern Microscopy Society (SEMS) is now online at the SEMS website in PDF format: http://www.biotech.ufl.edu/sems/
It contains information on the upcoming meeting in Clemson, SC. If you are a member, and have not received this notice via e-mail, and wish to be informed about the society through e-mail, please respond off-listserve at: jshields-at-cb.uga.edu
John Shields Center for Ultrastructural Research Univ. of Georgia Athens, GA
Such an instrument could provide for low-angle rotary shadowing capability for visualizing purified proteins at high resolution. It would need to have an electron beam gun for platinum evaporation and a separate one for carbon coating.
If this is what you have in mind, I support it. I purchased such an instrument that had good performance for about $20,000 about 10 years ago.
Dr. Joseph C. Besharse Professor and Chairman Dept of Cell Biology, Neurobiology and Anatomy Medical College of Wisconsin 8701 Watertown Plank Road Milwaukee, WI 53226-0509
} From: Gang Ning {gning-at-mcw.edu} } Organization: Medical College of Wisconsin } Reply-To: gning-at-mcw.edu } Date: Tue, 10 Apr 2001 14:17:42 -0500 } To: Microscopy Newsgroup {Microscopy-at-sparc5.microscopy.com} } Cc: "Dr. Traktman" {ptrakt-at-post.its.mcw.edu} , Ming Lei {mlei-at-mcw.edu} , "Dr. } Besharse" {jbeshars-at-mcw.edu} } Subject: Sputter coater } } Hi All: } } I want to buy a new/used sputter coater which enables to do rotary } shadowing as well as carbon coating. Any suggestions/input are } appreciated. } } Greg Ning } } EM Facility } Medical College of Wisconsin }
Question: Hi, I have just purchased a microscope, a good biological one. I need a microscope kit but nobody sells them around here. Anyway already I have glass slides and covers. I have read some on microscopy and I need an adhesive, resin I think its called to prepare slides? is this true? Also some ink to stain specimens. What are the names of all these chemicals so I can buy them all seperatly since no one sells them all together. Thankyou.
Dear Theresa: Although a metallographic laboratory, we use alcohol for specimen preparation, mainly for preparation of etchants and for specimen cleaning. We found the paperwork associated with pure ethanol onerous and switched to denatured alcohol Type 3A with no discernable difference. Beware of some of the denaturants, they produce unusual side effects.
Sam Purdy National Steel Tech Center Trenton MI
} ---------- } From: BOES,TERESA (HP-Corvallis,ex1) } Sent: Monday, April 9, 2001 2:50 PM } To: 'microscopy-at-MSA.microscopy.com' } Subject: Substitutes for absolute ethanol? } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } Has anyone used any of the denatured ethanols as a substitute for absolute } ethanol? } } We have recently run into some difficulty when needing to reorder 200 } proof } ethanol, which we use for dehydration and infiltration of samples } (primarily } many types of paper) prior to embedding in Spurrs epoxy, and for cleaning } samples (non paper) and lenses of light microscopes. The chemical company } selling the ethanol is insisting that we must have a liquor license before } they will ship to us. } } Ethanol denatured with a variety of substances is readily available and } can } be shipped with no licensing requirements. Our concern is that the } denaturing agent will leave a detectable residue on lenses, samples, and } may } cause problems with the polymerization of Spurrs. Rather than obtaining a } liquor license, we are considering using one of the 100:5 ethanol: } methanol } blends. If any of you have had successful or unsuccessful experiences } substituting denatured ethanol for absolute in embedding or cleaning } protocols, I would appreciate hearing from you. } } Teresa Boes } Hewlett-Packard } Analytical and Development Lab } 1000 Circle Blvd } Corvallis, OR 97330 } 541-715-7055 } teresa_boes-at-hp.com } }
I would caution biased opinions from installation engineer and sales representatives ie... James Fotinopoulos.... www.semguy.com...
Hmmmmmmmmm?
Food for thought
----- Original Message -----
} From: "jeanross" {jeanross-at-blue.weeg.uiowa.edu} } To: "Microscopy Listserver" {Microscopy-at-sparc5.microscopy.com} } Sent: Tuesday, April 10, 2001 10:22 AM } Subject: EDS summary } } } } ------------------------------------------------------------------------ } } 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 put together a summary of responses I got from my inquiry about a } week } } or so ago about EDS systems. I really appreciate everyone's input. We } } haven't made any decisions yet since we are still gathering information } but } } your responses will help. I've included the responses in their entirety } so I } } hope this helps others as well. } } } } Thanks again from everyone who contributed. } } } } Jean Ross } } Central Microscopy Research Facility } } University of Iowa } } } } -------------------------------------------------------------------------- } --- } } I have been using an IXRF EDS / Gresham Detector (with IXRF digital pulse } } processor) system for almost 2 years. Initially, the IXRF was installed on } } an ETEC. About 15 months ago the Etec was replaced with a Hitachi 3500, a } } new detector (Gresham) was purchased, and the EDS was installed on the } 3500. } } Generally, I am quite satisfied. There are minor software bugs, but IXRF } } has been reasonably good at fixing them when discovered. It has been my } } experience that all the systems have bugs, perhaps some more than others. } } Prior to the IXRF, I had a Kevex 8000/Delta. } } } } Low end noise and broad peaks were evident on first installation, but were } } soon fixed by tweaking the detector preamp and pulse processor amp. } } } } I am still running their first software package, "Iridium". I have the } } newest release, "EDS 2000", but lack of time has kept me from installing } and } } checking it out. } } } } I should mention that IXRF is a "virtual" company, with people spread out } } between Texas, California, etc. This has not proven to be a problem. } } } } Woody White } } McDermott Technology, Inc. } } nwwhite-at-mcdermott.com } } } } -------------------------------------------------------------------------- } --- } } } } } } We have had EDAX for about a decade and a half, and we are very pleased } with } } the product and the service we get. } } Carol Heckman } } heckman-at-bgnet.bgsu.edu } } } } -------------------------------------------------------------------------- } --- } } } } } } look at IXRF eds systems there web site is www.ixrfsystems.com, they are } } very affordable and offer no nonsense performance that second to none. } } } } happy ixrf user, } } } } James Fotinopoulos } } yzfrjim-at-ix.netcom.com } } } } } } -------------------------------------------------------------------------- } ---- } } } } I would recommend you give consideration to Doug Connors at } } TN Analyzer Service, Inc. of Dane, WI. Doug has rebuild and } } upgraded detectors for me for the last 6 years. He is dependable } } knowledgeable, and economical as well. } } } } Bob Roberts } } EM Lab Services, Inc. } } 2409 S. Rural Rd Suite C } } Tempe, Arizona 85282 } } (480) 967-3946 } } bobrobs-at-earthlink.net } } } } -------------------------------------------------------------------------- } --- } } } } } } We recently purchased a Noran Instruments Vantage DS1. We purchased this } } based on some very impressive demonstrations of software that the } salesperson } } brought in. Unfortunately they are still working out the bugs in their } } software. Everything they have is ported over from Unix, and literally } runs } } in a unix shell on an Microsoft Windows NT platform. This makes their } } software fairly buggy. Their response time to fix major bugs and hang-ups } in } } the software has been very slow, and if given the opportunity to do it all } } over again I'd probably look at Oxford Instruments. I would still rate } the } } quality of the equipment very high. Our detector performs at the } specified } } resolution, and is a good piece of equipment. Now if they could only get } the } } software end of it straight... } } My vote: } } 1) Oxford Instruments } } 2) Noran Instruments } } 3) Edax or some of the smaller players } } The benefit with going with a larger company is support and upgrades. We } have } } a 10 year old WDX that we just purchased new software and interface for } last } } year. Our old EDS was given some trade in value by Noran. And we all } know } } how valuable a service contract can be... } } } } Get back to me if you have any more questions, } } ~Jonathan } } Jonathan Dunlap } } Analytical Laboratory Manager } } Osram Sylvania Inc. } } 816 Lexington Avenue } } Warren, PA 16365 } } Ph: 814-726-6991 } } Fax: 814-726-6942 } } Jonathan.Dunlap-at-sylvania.com } } } } } } -------------------------------------------------------------------------- } ---- } } } } } } We have an Oxford Instruments Link ISIS Model 200 on our 2460N. We have } been } } happy with it, but I don't know about the direction that Oxford is } heading. I } } don't care for the feel of their new INCA software. Some might like it. It } } also seems to be slow coming together. Some of the functions are still } lacking } } after 2 (or is it 3) years of seeing it at MSA. } } } } We have an IXRF system on our JEOL 840A. It was a good price ($30K) for an } } upgrade to our Kevex several years ago. It does what we need. They keep at } } work on the software and have it freely available on the web. I might have } to } } pay closer attention and stay away from the beta stuff. They are still } working } } on it. They also have a nice digital pulse processor which stills stand } alone } } for about $5k. } } } } I still feel funny about some contacts with EVEX. I can't say much about } EDAX, } } NORAN, or PGT. They should all have good stuff but it might be pricey. The } } last we seriously looked at them was 6 years ago or so when we opted for } the } } Oxford. } } } } I was intrigued by the unit from Quartz PCI. I think it was called X-ray } One, } } or such. It was new at MSA 1-1/2 years ago but looked promising. } } } } Feel free to call if you want more details. } } } } Warren E Straszheim } } wesaia-at-iastate.edu } } } } -------------------------------------------------------------------------- } --- } } } } } } We purchased an EDAX Falcon system for our Hitachi S-3000N and I've been } } pleased with it. It has better light element sensitivity than most which } } was very important to me although I don't think that its mapping } } capabilities are as good as PGT's, say. I don't have direct experience } with } } Noran although I did talk to them and their system seemed ok - but } logistics } } didn't favor Noran so I passed on them. EDAX does have good integration } } with the Hitachi and the Quartz database. } } } } I'd be glad to respond more specifically if you'd like. } } } } Richard Shalvoy } } Arch Chemicals } } Cheshire, CT } } RBShalvoy-at-archchemicals.com } } } } -------------------------------------------------------------------------- } --- } } } } } } I have an iXRF systems out of Texas using a Gresham detector. It works } } well. Not the most cutting edge, but they are one of the "start ups". They } } have been around for I guess 6-7 years. I have a digital pulse processor } } and completely active control for x-ray maps and such. They are very price } } competitive, but lack a dedicated technical support person. You talk to } the } } programmer or electronics expert, but no techs on the phone whenever you } } have a software question. But, if you willing to wait a day for some } } answers then they are worth it. I haven't run across the problem where I } } thought, "if I just had a better system". If you want to integrated w/ WDS } } than maybe Noran. Also, if you want to integrated w/ motorized stage } } control, I don't think they off such a package, like the bigger companies. } } } } I have a Hitachi 450. I used to run a 2400 and 500 before I quit my day } job } } and went out on my own. I am very happy w/ Hitachi. } } } } Good Luck } } } } Fell free to call with any specifics. } } } } Their web page is www.ixrfsystems.com } } } } Ric } } } } SMARTech } } 860-491-3299 } } www.semguy.com } } 19 Cornwall Drive } } Goshen CT 06756 } } smartech-at-javanet.com } } } } -------------------------------------------------------------------------- } ---- } } } } } } Hello, all: } } } } I use Oxford ISIS300 system on HITACHI S-3500N (with VP mode) for light } } element analysis, mostly C, O, N, F, P, S, Si, Mg, as well as metal Co, } Ni-P, } } Pt, Cr, Fe, W, etc. This system works well. One useful function is the } overlay } } of 2 spectrums. I can easily subtract the blank from the sample spot and } make } } it easy to identify what is (are) in the sample. I am sure some other } program } } may have this kind of function, but I have not seen. } } } } Zhiyu Wang } } zhiyuw-at-home.com } } I would be interested in seeing the responses as I am going to try and get } } funding next year for a replacement for our EDAX PV9100 on an Hitachi } s-450. } } } } Dave } } David.Patton-at-uwe.ac.uk } } } } -------------------------------------------------------------------------- } --- } } } } } } We have been running EDX on SEMs and TEMs for many years. We used to have } a } } range of systems from Kevex, PGT, Noran, Link, EDAX, however a few years } ago } } we decided that we ought to standardize on one common system. After } evaluation } } we bought three Oxford Instruments ISIS systems. Whenever we have upgraded } or } } bought new systems they have been Oxford Instruments ISIS or now INCA. } } } } I have been happy with the ISIS except for the file handling that was not } } designed for a multi user facility such as ours (approx 120 EM users in } total } } roughly 25 to 30 swapping every year). I am really quite impressed by the } } INCA, Oxford Instruments are, at last, listening to the users and adding } user } } requested facilities. They have sorted out the file handling mess of the } ISIS } } and structured it well for an SEM user (not quite as well for a TEM user } but } } there are less of us). The software structure is quite intuitive and there } is } } a really impressive help menu and explanation of everything from the } physics } } of X-ray generation, how EM's work, how detectors work and how to analyze } } samples. } } } } Their detectors have always been good and the SATW (thin window detectors) } } still have a reasonable efficiency at low Z. B is possible but C is easy } and } } even the N peak is over 30% efficient (there is often a high absorption at } N). } } } } Another feature that is invaluable for TEM is the integral shutter that } will } } close when the count rate is too high. This protects the crystal, it } prevents } } it overloading and shutting down or worse the crystal efficiency may } change } } for a few minutes until it recovers fully. This may affects your } quantitative } } work. In TEM this is usually caused by hitting the grid bar and not really } a } } problem in SEM but I don't know what secondaries and ions you will have in } a } } variable pressure SEM. It could be useful for you, check with other high } } pressure SEM users. } } } } Regards, } } Ron } } } } Please note: Oxford Instruments have upgraded an ISIS to an INCA system in } my } } department, without charge, in return for access to the instrument for } } development projects and demonstrations for a fixed number of days. I } receive } } no benefit from this and the department has no benefit from Oxford } Instruments } } sales. I remain a thorn in the flesh of all our suppliers if I think they } } could improve their products or service. } } ron.doole-at-materials.oxford.ac.uk } } } } } } -------------------------------------------------------------------------- } ---- } } } } } } Hello, } } I am very familiar with the Oxford ISIS 300 series spectrometers. They are } } ok, and the new Inca system looks good too. However, I recently saw the } PGT } } spectrometer at Lehigh and it is very impressive. } } } } Steve } } Stephen_Skirius-at-bkitech.com } } } } } } } } -------------------------------------------------------------------------- } } } } } } I'm also in the market for an EDS system and have looked at EDAX, Noran, } } PGT and Oxford. } } } } I edited out PGT because in order to quantify you have to optimize the } } system for the type of sample by playing with fudge factors, which none of } } the other systems have to do (though one of them, I think Noran, lets you } } adjust a sensitivity factor if you want to, but they didn't do it on my } } samples that were tested against known microprobe results and the answers } } were fine). I also eliminated Oxford, though it has a terrific user } } interface (maybe at the expense of functionality), because they } } consistently IDed my aluminum peaks as Br or Tm (!); this made me wonder } } about all their algorithms. They claim it had to do with the takeoff angle } } on the particular SEM being used, but that shouldn't be a factor. } } } } I like both EDAX and Noran, though for different reasons. EDAX user } } interface is better than Noran's, though again, I think Noran possibly } } offers more routines (it's hard to keep track and see absolutely } everything } } a system has to offer in a demo day....).Noran can multitask - work on } } several programs while a spectral map is being collected, for instance } } (does EDAX? I have to check). But EDAX has a beam skirt reduction routine } } for low vac mode (though it's time consuming, so a bit cumbersome), and } } their peak modeling is right up front - but Noran can put theirs up front } } also if you want to have it accessible (yes) and I think Noran might be a } } little better engineered. } } } } As you can see I'm still in a quandary (ditto for the two contending SEMs, } } LEO and ESEM). Whatever I decide I'll still be very interested in the } } results of your posting - especially if other folks' info comes in within } } the next week or so it would help in my decision too. } } } } I hope my input helps a little. Good luck with your quest! } } } } Dee Breger } } micro-at-ldeo.columbia.edu } } } } } } -------------------------------------------------------------------------- } - } } } } } } I looked into Noran, EDAX, and PGT. Noran was quickly culled (less user } } friendly, less abilities, didn't work right during demo), but EDAX and PGT } } both seemed to have equivalent capabilities (PGT claimed a 'proprietary' } } signal amplifier/digitizer doohickey, but it was only in the placement.) } } For the long-time spectrum gathering (I forget the technical term), PGT } } makes many passes with short dwell times while EDAX dwells on each pixel } } much longer to collect data & does it in 1 pass. Kinda 6 of one, half } dozen } } of the other. What made us choose the EDAX Phoenix system was the fact } that } } the PGT software was UNIX-based (although hidden) while EDAX is PC-based. } } I've heard rumors that PGT is switching to PC-based; we purchased our } system } } in 1999. I've also heard that Noran has practically no service techs (but } } that may be an East coast thing.) We've been happy with the EDAX service, } } and I enjoyed their user school very much. By the way, our SEM is a } } variable pressure JEOL 5900, and it's integrated with the EDAX system. } } } } Hope I've been helpful, } } } } Jane L. LaGoy } } Development Engineer } } Bodycote IMT, Inc. } } 155 River Street } } Andover, MA 01810 } } 978-470-1620 } } jlagoy-at-bodycote-imt.com } } } } } } } } } } }
{HTML} {FONT FACE=arial,helvetica} Jean, {BR} {BR} I would caution biased opinions from installation engineer and sales {BR} representatives ie... James Fotinopoulos.... www.semguy.com... {BR} {BR} Hmmmmmmmmm? {BR} {BR} Food for thought {BR} {FONT SIZE=2} {BR} {BR} {BR} {BR} {BR} ----- Original Message ----- {BR} {/FONT} {FONT COLOR="#000000" SIZE=3 FAMILY="SANSSERIF" FACE="Arial" LANG="0"} {BR} {/FONT} {FONT COLOR="#000000" SIZE=2 FAMILY="SANSSERIF" FACE="Arial" LANG="0"} {BLOCKQUOTE TYPE=CITE style="BORDER-LEFT: #0000ff 2px solid; MARGIN-LEFT: 5px; MARGIN-RIGHT: 0px; PADDING-LEFT: 5px"} From: "jeanross" <jeanross-at-blue.weeg.uiowa.edu> {BR} To: "Microscopy Listserver" <Microscopy-at-sparc5.microscopy.com> {BR} Sent: Tuesday, April 10, 2001 10:22 AM {BR} Subject: EDS summary {BR} {BR} {BR} > ------------------------------------------------------------------------ {BR} > The Microscopy ListServer -- Sponsor: The Microscopy Society of America {BR} > To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com {BR} > On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html {BR} > -----------------------------------------------------------------------. {BR} > {BR} > {BR} > I have put together a summary of responses I got from my inquiry about a {BR} week {BR} > or so ago about EDS systems. I really appreciate everyone's input. We {BR} > haven't made any decisions yet since we are still gathering information {BR} but {BR} > your responses will help. I've included the responses in their entirety {BR} so I {BR} > hope this helps others as well. {BR} > {BR} > Thanks again from everyone who contributed. {BR} > {BR} > Jean Ross {BR} > Central Microscopy Research Facility {BR} > University of Iowa {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > I have been using an IXRF EDS / Gresham Detector (with IXRF digital pulse {BR} > processor) system for almost 2 years. Initially, the IXRF was installed on {BR} > an ETEC. About 15 months ago the Etec was replaced with a Hitachi 3500, a {BR} > new detector (Gresham) was purchased, and the EDS was installed on the {BR} 3500. {BR} > Generally, I am quite satisfied. There are minor software bugs, but IXRF {BR} > has been reasonably good at fixing them when discovered. It has been my {BR} > experience that all the systems have bugs, perhaps some more than others. {BR} > Prior to the IXRF, I had a Kevex 8000/Delta. {BR} > {BR} > Low end noise and broad peaks were evident on first installation, but were {BR} > soon fixed by tweaking the detector preamp and pulse processor amp. {BR} > {BR} > I am still running their first software package, "Iridium". I have the {BR} > newest release, "EDS 2000", but lack of time has kept me from installing {BR} and {BR} > checking it out. {BR} > {BR} > I should mention that IXRF is a "virtual" company, with people spread out {BR} > between Texas, California, etc. This has not proven to be a problem. {BR} > {BR} > Woody White {BR} > McDermott Technology, Inc. {BR} > nwwhite-at-mcdermott.com {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > {BR} > {BR} > We have had EDAX for about a decade and a half, and we are very pleased {BR} with {BR} > the product and the service we get. {BR} > Carol Heckman {BR} > heckman-at-bgnet.bgsu.edu {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > {BR} > {BR} > look at IXRF eds systems there web site is www.ixrfsystems.com, they are {BR} > very affordable and offer no nonsense performance that second to none. {BR} > {BR} > happy ixrf user, {BR} > {BR} > James Fotinopoulos {BR} > yzfrjim-at-ix.netcom.com {BR} > {BR} > {BR} > -------------------------------------------------------------------------- {BR} ---- {BR} > {BR} > I would recommend you give consideration to Doug Connors at {BR} > TN Analyzer Service, Inc. of Dane, WI. Doug has rebuild and {BR} > upgraded detectors for me for the last 6 years. He is dependable {BR} > knowledgeable, and economical as well. {BR} > {BR} > Bob Roberts {BR} > EM Lab Services, Inc. {BR} > 2409 S. Rural Rd Suite C {BR} > Tempe, Arizona 85282 {BR} > (480) 967-3946 {BR} > bobrobs-at-earthlink.net {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > {BR} > {BR} > We recently purchased a Noran Instruments Vantage DS1. We purchased this {BR} > based on some very impressive demonstrations of software that the {BR} salesperson {BR} > brought in. Unfortunately they are still working out the bugs in their {BR} > software. Everything they have is ported over from Unix, and literally {BR} runs {BR} > in a unix shell on an Microsoft Windows NT platform. This makes their {BR} > software fairly buggy. Their response time to fix major bugs and hang-ups {BR} in {BR} > the software has been very slow, and if given the opportunity to do it all {BR} > over again I'd probably look at Oxford Instruments. I would still rate {BR} the {BR} > quality of the equipment very high. Our detector performs at the {BR} specified {BR} > resolution, and is a good piece of equipment. Now if they could only get {BR} the {BR} > software end of it straight... {BR} > My vote: {BR} > 1) Oxford Instruments {BR} > 2) Noran Instruments {BR} > 3) Edax or some of the smaller players {BR} > The benefit with going with a larger company is support and upgrades. We {BR} have {BR} > a 10 year old WDX that we just purchased new software and interface for {BR} last {BR} > year. Our old EDS was given some trade in value by Noran. And we all {BR} know {BR} > how valuable a service contract can be... {BR} > {BR} > Get back to me if you have any more questions, {BR} > ~Jonathan {BR} > Jonathan Dunlap {BR} > Analytical Laboratory Manager {BR} > Osram Sylvania Inc. {BR} > 816 Lexington Avenue {BR} > Warren, PA 16365 {BR} > Ph: 814-726-6991 {BR} > Fax: 814-726-6942 {BR} > Jonathan.Dunlap-at-sylvania.com {BR} > {BR} > {BR} > -------------------------------------------------------------------------- {BR} ---- {BR} > {BR} > {BR} > We have an Oxford Instruments Link ISIS Model 200 on our 2460N. We have {BR} been {BR} > happy with it, but I don't know about the direction that Oxford is {BR} heading. I {BR} > don't care for the feel of their new INCA software. Some might like it. It {BR} > also seems to be slow coming together. Some of the functions are still {BR} lacking {BR} > after 2 (or is it 3) years of seeing it at MSA. {BR} > {BR} > We have an IXRF system on our JEOL 840A. It was a good price ($30K) for an {BR} > upgrade to our Kevex several years ago. It does what we need. They keep at {BR} > work on the software and have it freely available on the web. I might have {BR} to {BR} > pay closer attention and stay away from the beta stuff. They are still {BR} working {BR} > on it. They also have a nice digital pulse processor which stills stand {BR} alone {BR} > for about $5k. {BR} > {BR} > I still feel funny about some contacts with EVEX. I can't say much about {BR} EDAX, {BR} > NORAN, or PGT. They should all have good stuff but it might be pricey. The {BR} > last we seriously looked at them was 6 years ago or so when we opted for {BR} the {BR} > Oxford. {BR} > {BR} > I was intrigued by the unit from Quartz PCI. I think it was called X-ray {BR} One, {BR} > or such. It was new at MSA 1-1/2 years ago but looked promising. {BR} > {BR} > Feel free to call if you want more details. {BR} > {BR} > Warren E Straszheim {BR} > wesaia-at-iastate.edu {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > {BR} > {BR} > We purchased an EDAX Falcon system for our Hitachi S-3000N and I've been {BR} > pleased with it. It has better light element sensitivity than most which {BR} > was very important to me although I don't think that its mapping {BR} > capabilities are as good as PGT's, say. I don't have direct experience {BR} with {BR} > Noran although I did talk to them and their system seemed ok - but {BR} logistics {BR} > didn't favor Noran so I passed on them. EDAX does have good integration {BR} > with the Hitachi and the Quartz database. {BR} > {BR} > I'd be glad to respond more specifically if you'd like. {BR} > {BR} > Richard Shalvoy {BR} > Arch Chemicals {BR} > Cheshire, CT {BR} > RBShalvoy-at-archchemicals.com {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > {BR} > {BR} > I have an iXRF systems out of Texas using a Gresham detector. It works {BR} > well. Not the most cutting edge, but they are one of the "start ups". They {BR} > have been around for I guess 6-7 years. I have a digital pulse processor {BR} > and completely active control for x-ray maps and such. They are very price {BR} > competitive, but lack a dedicated technical support person. You talk to {BR} the {BR} > programmer or electronics expert, but no techs on the phone whenever you {BR} > have a software question. But, if you willing to wait a day for some {BR} > answers then they are worth it. I haven't run across the problem where I {BR} > thought, "if I just had a better system". If you want to integrated w/ WDS {BR} > than maybe Noran. Also, if you want to integrated w/ motorized stage {BR} > control, I don't think they off such a package, like the bigger companies. {BR} > {BR} > I have a Hitachi 450. I used to run a 2400 and 500 before I quit my day {BR} job {BR} > and went out on my own. I am very happy w/ Hitachi. {BR} > {BR} > Good Luck {BR} > {BR} > Fell free to call with any specifics. {BR} > {BR} > Their web page is www.ixrfsystems.com {BR} > {BR} > Ric {BR} > {BR} > SMARTech {BR} > 860-491-3299 {BR} > www.semguy.com {BR} > 19 Cornwall Drive {BR} > Goshen CT 06756 {BR} > smartech-at-javanet.com {BR} > {BR} > -------------------------------------------------------------------------- {BR} ---- {BR} > {BR} > {BR} > Hello, all: {BR} > {BR} > I use Oxford ISIS300 system on HITACHI S-3500N (with VP mode) for light {BR} > element analysis, mostly C, O, N, F, P, S, Si, Mg, as well as metal Co, {BR} Ni-P, {BR} > Pt, Cr, Fe, W, etc. This system works well. One useful function is the {BR} overlay {BR} > of 2 spectrums. I can easily subtract the blank from the sample spot and {BR} make {BR} > it easy to identify what is (are) in the sample. I am sure some other {BR} program {BR} > may have this kind of function, but I have not seen. {BR} > {BR} > Zhiyu Wang {BR} > zhiyuw-at-home.com {BR} > I would be interested in seeing the responses as I am going to try and get {BR} > funding next year for a replacement for our EDAX PV9100 on an Hitachi {BR} s-450. {BR} > {BR} > Dave {BR} > David.Patton-at-uwe.ac.uk {BR} > {BR} > -------------------------------------------------------------------------- {BR} --- {BR} > {BR} > {BR} > We have been running EDX on SEMs and TEMs for many years. We used to have {BR} a {BR} > range of systems from Kevex, PGT, Noran, Link, EDAX, however a few years {BR} ago {BR} > we decided that we ought to standardize on one common system. After {BR} evaluation {BR} > we bought three Oxford Instruments ISIS systems. Whenever we have upgraded {BR} or {BR} > bought new systems they have been Oxford Instruments ISIS or now INCA. {BR} > {BR} > I have been happy with the ISIS except for the file handling that was not {BR} > designed for a multi user facility such as ours (approx 120 EM users in {BR} total {BR} > roughly 25 to 30 swapping every year). I am really quite impressed by the {BR} > INCA, Oxford Instruments are, at last, listening to the users and adding {BR} user {BR} > requested facilities. They have sorted out the file handling mess of the {BR} ISIS {BR} > and structured it well for an SEM user (not quite as well for a TEM user {BR} but {BR} > there are less of us). The software structure is quite intuitive and there {BR} is {BR} > a really impressive help menu and explanation of everything from the {BR} physics {BR} > of X-ray generation, how EM's work, how detectors work and how to analyze {BR} > samples. {BR} > {BR} > Their detectors have always been good and the SATW (thin window detectors) {BR} > still have a reasonable efficiency at low Z. B is possible but C is easy {BR} and {BR} > even the N peak is over 30% efficient (there is often a high absorption at {BR} N). {BR} > {BR} > Another feature that is invaluable for TEM is the integral shutter that {BR} will {BR} > close when the count rate is too high. This protects the crystal, it {BR} prevents {BR} > it overloading and shutting down or worse the crystal efficiency may {BR} change {BR} > for a few minutes until it recovers fully. This may affects your {BR} quantitative {BR} > work. In TEM this is usually caused by hitting the grid bar and not really {BR} a {BR} > problem in SEM but I don't know what secondaries and ions you will have in {BR} a {BR} > variable pressure SEM. It could be useful for you, check with other high {BR} > pressure SEM users. {BR} > {BR} > Regards, {BR} > Ron {BR} > {BR} > Please note: Oxford Instruments have upgraded an ISIS to an INCA system in {BR} my {BR} > department, without charge, in return for access to the instrument for {BR} > development projects and demonstrations for a fixed number of days. I {BR} receive {BR} > no benefit from this and the department has no benefit from Oxford {BR} Instruments {BR} > sales. I remain a thorn in the flesh of all our suppliers if I think they {BR} > could improve their products or service. {BR} > ron.doole-at-materials.oxford.ac.uk {BR} > {BR} > {BR} > -------------------------------------------------------------------------- {BR} ---- {BR} > {BR} > {BR} > Hello, {BR} > I am very familiar with the Oxford ISIS 300 series spectrometers. They are {BR} > ok, and the new Inca system looks good too. However, I recently saw the {BR} PGT {BR} > spectrometer at Lehigh and it is very impressive. {BR} > {BR} > Steve {BR} > Stephen_Skirius-at-bkitech.com {BR} > {BR} > {BR} > {BR} > -------------------------------------------------------------------------- {BR} > {BR} > {BR} > I'm also in the market for an EDS system and have looked at EDAX, Noran, {BR} > PGT and Oxford. {BR} > {BR} > I edited out PGT because in order to quantify you have to optimize the {BR} > system for the type of sample by playing with fudge factors, which none of {BR} > the other systems have to do (though one of them, I think Noran, lets you {BR} > adjust a sensitivity factor if you want to, but they didn't do it on my {BR} > samples that were tested against known microprobe results and the answers {BR} > were fine). I also eliminated Oxford, though it has a terrific user {BR} > interface (maybe at the expense of functionality), because they {BR} > consistently IDed my aluminum peaks as Br or Tm (!); this made me wonder {BR} > about all their algorithms. They claim it had to do with the takeoff angle {BR} > on the particular SEM being used, but that shouldn't be a factor. {BR} > {BR} > I like both EDAX and Noran, though for different reasons. EDAX user {BR} > interface is better than Noran's, though again, I think Noran possibly {BR} > offers more routines (it's hard to keep track and see absolutely {BR} everything {BR} > a system has to offer in a demo day....).Noran can multitask - work on {BR} > several programs while a spectral map is being collected, for instance {BR} > (does EDAX? I have to check). But EDAX has a beam skirt reduction routine {BR} > for low vac mode (though it's time consuming, so a bit cumbersome), and {BR} > their peak modeling is right up front - but Noran can put theirs up front {BR} > also if you want to have it accessible (yes) and I think Noran might be a {BR} > little better engineered. {BR} > {BR} > As you can see I'm still in a quandary (ditto for the two contending SEMs, {BR} > LEO and ESEM). Whatever I decide I'll still be very interested in the {BR} > results of your posting - especially if other folks' info comes in within {BR} > the next week or so it would help in my decision too. {BR} > {BR} > I hope my input helps a little. Good luck with your quest! {BR} > {BR} > Dee Breger {BR} > micro-at-ldeo.columbia.edu {BR} > {BR} > {BR} > -------------------------------------------------------------------------- {BR} - {BR} > {BR} > {BR} > I looked into Noran, EDAX, and PGT. Noran was quickly culled (less user {BR} > friendly, less abilities, didn't work right during demo), but EDAX and PGT {BR} > both seemed to have equivalent capabilities (PGT claimed a 'proprietary' {BR} > signal amplifier/digitizer doohickey, but it was only in the placement.) {BR} > For the long-time spectrum gathering (I forget the technical term), PGT {BR} > makes many passes with short dwell times while EDAX dwells on each pixel {BR} > much longer to collect data & does it in 1 pass. Kinda 6 of one, half {BR} dozen {BR} > of the other. What made us choose the EDAX Phoenix system was the fact {BR} that {BR} > the PGT software was UNIX-based (although hidden) while EDAX is PC-based. {BR} > I've heard rumors that PGT is switching to PC-based; we purchased our {BR} system {BR} > in 1999. I've also heard that Noran has practically no service techs (but {BR} > that may be an East coast thing.) We've been happy with the EDAX service, {BR} > and I enjoyed their user school very much. By the way, our SEM is a {BR} > variable pressure JEOL 5900, and it's integrated with the EDAX system. {BR} > {BR} > Hope I've been helpful, {BR} > {BR} > Jane L. LaGoy {BR} > Development Engineer {BR} > Bodycote IMT, Inc. {BR} > 155 River Street {BR} > Andover, MA 01810 {BR} > 978-470-1620 {BR} > jlagoy-at-bodycote-imt.com {BR} > {BR} > {BR} > {BR} > {BR} {BR} {/XMP} {/FONT} {FONT COLOR="#0f0f0f" SIZE=2 FAMILY="SANSSERIF" FACE="Arial" LANG="0"} {BR} {/FONT} {/HTML}
Return-Path: {ptarq-at-evex.com} Received: from rly-xb03.mx.aol.com (rly-xb03.mail.aol.com [172.20.105.104]) by air-xb02.mail.aol.com (v77_r1.36) with ESMTP; Tue, 10 Apr 2001 17:56:34 -0500 Received: from mxr01.nyc01.dsl.net (mxr01.nyc01.dsl.net [216.175.203.53]) by rly-xb03.mx.aol.com (v77_r1.36) with ESMTP; Tue, 10 Apr 2001 17:56:05 -0400 Received: from pb133nt (64-51-105-68.client.dsl.net [64.51.105.68]) by mxr01.nyc01.dsl.net (Postfix) with SMTP id 1F49C34538 for {lhconsulting-at-aol.com} ; Tue, 10 Apr 2001 17:52:57 -0400 (EDT) Message-ID: {002b01c0c208$95dfe860$6601a8c0-at-evex.com} Reply-To: "Evex Mail" {ptarq-at-evex.com} } From: "Evex Mail" {ptarq-at-evex.com} To: {lhconsulting-at-aol.com}
I guess it was a slip: In a sputter coater you can do rotary coating but shadowing is possible in evaporators only. For carbon coating the sputter coater would need an attachment for carbon string evaporation, which may be used for SEM and analyses, but not in TEM. Disclaimer: ProSciTech is the EMITECH instrument distributor in Australasia. 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 ABN: 99 724 136 560 www.proscitech.com
On Wednesday, April 11, 2001 5:18 AM, Gang Ning [SMTP:gning-at-mcw.edu] wrote: } } } Hi All: } } I want to buy a new/used sputter coater which enables to do rotary } shadowing as well as carbon coating. Any suggestions/input are } appreciated. } } Greg Ning } } EM Facility } Medical College of Wisconsin }
A colleague and I each recently bought Microtek scanners to scan TEM negatives. I have the Artixscan 1100 and he has the Model 8700 which has similar characteristics (actually higher resolution -1200dpi), 3.9 dmax at 42 bits color (14 grayscale), and the glassless film carrier setup. The 8700 has USB and Firewire interfaces and is cheaper ( {$1000), and the 1000 dpi Model 1100 has a SCSI interface. You might want to check out the specs of the lower cost model 8700 on the microtekusa website if your computer can handle USB or Firewire. Both scanners have performed up to our expectations, which I would characterize as modest. Microtek does not supply a 3-1/4 x 4 " negative carrier for standard size TEM film but you can easily make a serviceable one from stiff paper or light cardboard.
How much scanner resolution should you buy? The answer depends on how you intend to use it. Most applications do not require capturing the full resolution of the negative. From a practical viewpoint, the scanner resolution just determines how many times you can magnify the negative image to produce the final print size. For example, to get a publication-size print at 300 dpi, an image scanned at 1200 dpi scan could be zoomed 4X. A practical alternative to spending more for higher scanning resolution is to take photos at higher magnification. One exception is with lattice images from the TEM, which (depending on the lattice fringe spacing on the negative) might require higher scan resolutions to avoid getting a moire effect. (Of course, not everyone agrees. My colleague prefers to always scan at the maximum resolution).
What does a Dmax of 3.9 mean to you? To me it means a very dark negative. D is the log of the transmitted to incident intensity ratio. I wonder if users ever actually verify the manufacturer's specs with a calibrated density target. A Dmax of 3.9 can be useful for scanning TEM diffraction patterns that might have high contrast, but TEM micrograph negatives of metals and ceramics generally don't have that much contrast and biological thin section photos tend to have rather weak contrast. If your negatives are simply dark, use shorter photo exposure times. Scanning with maximum allowed grayscale resolutions (e.g., 14 bits rather than 8) is highly recommended if you intend to enhance or adjust images, but that's another story.
Larry Thomas Pacific Northwest National Laboratory MSIN P8-16 P.O. Box 999 Richland, WA 99352 Phone: (509)372-0793 Fax: (509)376-6308 Email: mailto:Larry.Thomas-at-pnl.gov
---------- From: Tom Phillips Sent: Tuesday, April 10, 2001 11:22 AM To: Alwyn Eades Cc: Microscopy-at-sparc5.microscopy.com Subject: Re: Scanners
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I too am about to buy and I would make a couple of comments on your evaluation. First, let me remind everyone that the Dynamic range is a log scale so small numerical differences are significant.
I also think the Nikon Coolscan 8000 looks great but it only takes a 2.5 x 3.5 negative which is smaller than my JEOL and Hitachi EM negative sizes (~ 3 1/2 by 4 1/2"). Have these EM manufacturers gone to a smaller film size or is Nikon using a non-Japanese EM as their standard? seems odd but I don't see how the Nikon would be very useful. You say a {2000 line scanner would be useful 9 out of 10 times but want the 2000+ lines for the occasional high res scan. I would argue that the size of the negative was the more important variable to be worried about. The Nikon couldn't handle 4x5 LM negatives or transparencies from autoradiography of Westerns/Northerns, etc.
My leading candidate is the ArtixScan 1100 has a Dmax of 3.9 (about $1600 with SCSI card). This was has a 1000 x 2000 dpi resolution. more details at www.microtek.com. This is my leading candidate. It was 4 negative carriers and I await word whether one could be modified to carry a 3 1/2 by 4 1/2 negative. At worst, I will have my scientific instrumentation shop guys fabricate a holder. It comes with a glass 8 x 10 glass carrier for odd size negs but I want to avoid Newton rings and want a glassless carrier.
I would appreciate comments on the following argument (I think I have this correctly figured out but am not sure since so many out there seem to want to have a higher resolution scanner). I have a Fuji Pictrography 3000 printer with a 400 dpi output that is as good as any other widely available printer in the academic world. If you figure the maximum published image size is about 8 inches, that would mean the maximum image size be 3200 dpi wide. A 1000 dpi scan of my negative would be 4500 x 3500 dpi. I could crop by about 28% or 10% depending on the orientation of the negative and still be taking full advantage of the printer resolution. In reality, most EM publication prints are smaller than 8" wide so one could crop even more and still not need more than 1000 dpi. A resolution } 1000 dpi would be useful for subtle morphometric analysis but a 4000 dpi scan of a 3 x 4 negative would be 192 MB. That is pretty big for doing morphometry on! A 1000 dpi scan of a 3.5 x 4.5" negative would be about 16 MB and that is much more manageable. Perhaps the difference is in the type of EM we are doing. I am working with biological specimens doing standard thin section type stuff. are you doing some Material Sci application that demands more?
I will be interested in Alwyn (and any others) reply since I hope to buy one soon!
} . } } } There was a thread recently on scanners for TEM film. I have looked up } all the models mentioned, on the web and called agents for prices - and } produced a comparative table, given below. } } I do not guarantee that the figures are accurate but they are my best } interpretation of the data given. } } In the light of experience and Nestor's comments, I would suggest that } 2000 dpi is a minimum for TEM negatives. You may be able to get away } with less nine times out of ten, but there will be occasions when you } need more. } I would exclude the Minolta and all the Epsons from consideration } (despite the incredibly low prices of some of the Epsons) because of the } low pixel density. } } Among the rest the Nikon has the best pixel density and the best optical } density (another critical parameter for TEM negatives). The price is } very competitive too. The Nikon web site does not give a time for } scanning a negative. On the face of it the Nikon would be a best buy - } get a separate, inexpensive flatbed scanner for the other work. } } These comments are all my own opinions based on manufacturers' data. } Since we are considering purchase any comments to the contrary would be } most welcome. } } } } Code Maker Model Type } } } } A Agfa DuoScan T2500 Flatbed } -Transparency included } } B Epson 1640 several versions Flatbed } -Transparency option } 1680 several versions } } C 1600 several versions Flatbed } -Transparency included } } D Imacon Flextight Precision II Drum -for } film and large format } } E Minolta Dimage ScanMulti II Film } } F Nikon Super Coolscan 8000ED Film } } G Polaroid 45 Ultra Film } } H Umax Powerlook 3000 Flatbed } -Transparency included } } } } } } Code dpi OD Time Price } Opinion } at 6 x 9 cm } } } A 2500 x2500 3.4 3 min } $4,500 Fair } } B 1600 x 3200 3.6 } $300-$3000 Poor } } $800-$1400 Poor } } C 1600 x 3200 3.3 } $650-$1160 Not suitable } } D 2240 x2240** 3.9/4.1 N/A above } $10k Good: low pixel density } } E 1128 x 1128 3.6 } Not suitable } } F 4000 x 4000 4.2 N/A } $2,695 V. Good } } G 2500 x 2500 3.8 5 min } $7,495 Good but pricey } } H 3048 x 3048 3.6 3 min } $6,499 Good } } } -- } .......... } Alwyn Eades } Department of Materials Science and Engineering } Lehigh University } 5 East Packer Avenue } Bethlehem } Pennsylvania 18015-3195 } Phone 610 758 4231 } Fax 610 758 4244 } jae5-at-lehigh.edu
-- 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)
Noise is only part of the problem, or solution, when considering a quantitative approach to scanning in negative or positive images. Since noise in electronic detection systems is largely dependent on temperature, that must also be brought into consideration. But the characteristics of the detector can be even more important. An array type device, such as a CCD, can have characteristics that vary from pixel to pixel as sensitivity, dynamic range and noise susceptibility. Your desire for a Consumer's Report on scanners is probably most appropriate as these various effects are difficult, if not impossible, to measure in a lab. Even if possible, these measurements may not be extendable to similar machines that aren't individually tested. Best to try to find a consensus on visual traits.
On Tuesday, April 10, 2001 7:50 AM, Sinkler, Wharton [SMTP:WSinkler-at-uop.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 } -----------------------------------------------------------------------. } } } } Laurie and Gary, } } Would noise be a good criterion? Say, for a perfectly evenly darkened film } (if such a thing existed, or at least even on a scale { { collected pixel } size) - what is the value of the noise (standard deviation of pixel value) } as a function of film darkness (density)? } } This would presumably improve with the time of collection. Thus how "good" } your scanner is depends on how you run it or whether it lets you take a } slower scan or to average multiple scans. With the exception of drum } scanners these devices all use CCD arrays. So what is probably most of } interest is the signal to noise ratio as a function of illumination } intensity, with everything known about CCD's going into determining this. } The maximum density the scanner can handle is just the point at which the } noise swamps the signal. } } There must be some good literature out there on the sources of noise, } optimizing collection (scan) time etc. One article which might be a } starting point is: } } G. H. Campbell, W. E. King and D. Cohen "Analysis of Experimental Error in } High Resolution Electron Micrographs", Microscopy and Microanalysis vol. 3 } (1997) p. 451. } } This is not very detailed, and treats only the total random noise, i.e. } grouping noise arising in collecting the image with that arising from the } scanner. } } Now, finding a good "Consumer Report" test with hard numbers on commercial } models is likely to be a lot harder! } } Wharton } } } -----Original Message----- } } From: Gary Gaugler [SMTP:gary-at-gaugler.com] } } Sent: Monday, April 09, 2001 10:37 PM } } To: L. D. Marks } } Cc: MSA listserver } } Subject: Re: Scanners: quantitative accuracy } } } } -------------------------------------------------------------------- ---- } } 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 } } -----------------------------------------------------------------------. } } } } } } How do you define "quantitative digitization?" i.e., what } } variables are you dealing with in this respect? What are } } the "absolute terms?" } } } } Anyone else have some ideas about this topic? } } } } gg } } } } } } At 10:10 AM 4/9/2001, you wrote: } } } } } I have been listening to the thread on scanners. Has anyone done } } } tests of how accurate they are in absolute terms for quantitative } } } digitization? } } } } } } ------------------------------------------------------- } } } Laurence Marks } } } Department of Materials Science and Engineering & } } } Center for Transportation Nanotechnology } } } Northwestern University } } } Tel: (847) 491-3996 Fax: (847) 491-7820 } } } mailto:ldm-at-risc4.numis.nwu.edu } } } http://www.numis.nwu.edu http://www.ctn.northwestern.edu } } } }
Allen R. Sampson, Owner Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174 voice 630.513.7093 fax 630.513.7092
Question: My student accidently got immersion oil on the 40x Leica Plan Acromat. I took a chance and used a fine artist's brush and xylene to clean it recalling (I may be wrong) that lens adhesives are soluble in alcohols not xylene, toluene and similar. Well, the lens didn't fall out. Too bad because I need an excuse to upgrade! If (or when) this happens again, what would you recommend for cleaning?
Does anyone out there have any experience microtoming bone that has NOT been decalcified? Is it even possible? I'm hoping to be able to have an answer for the admin people--prior to simply trying it out and possibly damaging my diamond knife.
Don Moravits Senior Technician Southwest Research Institute 6220 Culebra Road San Antonio, Texas 78238
I am embedding feather barbs which are mostly spongey dry collagen with keratin and small air pockets.
I am attempting Jan Dycks method-
1. 0.25 M NaOH 30 mins 2. formic acid / absolute alcohol 2:3 2 hrs ( to fill air pockets with solution) 3. 15% epon / propylene oxide 3 days 4. standard graduated increase of epon / dehydrant
Any other suggestions? Possibly from other similar material such as plant material.
Thanks
Tim Quinn Kansas University Museum of Natural History Lawrence, KS 785-864-4556
I would love to take advantage of the Firewire option but my information is that the 8700 has a Dmax of 3.4 vs the 3.9 for the 1100. That is a significant difference. Do EM negatives of biological thin sections reach that? I think so. I do a lot of EM immunocytochemistry and have to look for gold (intensely black) against a very dark tissue component so I am hoping the higher Dmax improves my results. I frequently scan negatives on a Umax 1100 (Dmax 3.4??) and can't differentiate the gold from the background although by eye I can discriminate them when the negative is placed on a light box. Changing my exposure would give me an unuseable image for the rest of the tissue. Maybe this is an extreme case but I suspect that lots of "dark organelles" (e.g., lysosomes, nuclei) have fine structure that get lost in the scanning with a low Dmax scanner. Tom
} A colleague and I each recently bought Microtek scanners to scan TEM } negatives. } I have the Artixscan 1100 and he has the Model 8700 which has similar } characteristics (actually higher resolution -1200dpi), 3.9 dmax at } 42 bits color } (14 grayscale), and the glassless film carrier setup. The 8700 has USB and } Firewire interfaces and is cheaper ( {$1000), and the 1000 dpi Model 1100 has a } SCSI interface. You might want to check out the specs of the lower cost model } 8700 on the microtekusa website if your computer can handle USB or Firewire. } Both scanners have performed up to our expectations, which I would } characterize } as modest. Microtek does not supply a 3-1/4 x 4 " negative carrier } for standard } size TEM film but you can easily make a serviceable one from stiff paper or } light cardboard. } } How much scanner resolution should you buy? The answer depends on how you } intend to use it. Most applications do not require capturing the full } resolution of the negative. From a practical viewpoint, the scanner } resolution } just determines how many times you can magnify the negative image to } produce the } final print size. For example, to get a publication-size print at 300 dpi, an } image scanned at 1200 dpi scan could be zoomed 4X. A practical alternative to } spending more for higher scanning resolution is to take photos at higher } magnification. One exception is with lattice images from the TEM, which } (depending on the lattice fringe spacing on the negative) might require higher } scan resolutions to avoid getting a moire effect. (Of course, not everyone } agrees. My colleague prefers to always scan at the maximum resolution). } } What does a Dmax of 3.9 mean to you? To me it means a very dark } negative. D is } the log of the transmitted to incident intensity ratio. I wonder if } users ever } actually verify the manufacturer's specs with a calibrated density target. A } Dmax of 3.9 can be useful for scanning TEM diffraction patterns that } might have } high contrast, but TEM micrograph negatives of metals and ceramics generally } don't have that much contrast and biological thin section photos tend to have } rather weak contrast. If your negatives are simply dark, use shorter photo } exposure times. Scanning with maximum allowed grayscale resolutions (e.g., 14 } bits rather than 8) is highly recommended if you intend to enhance or adjust } images, but that's another story. } } } Larry Thomas } Pacific Northwest National Laboratory } MSIN P8-16 } P.O. Box 999 } Richland, WA 99352 } Phone: (509)372-0793 Fax: (509)376-6308 } Email: mailto:Larry.Thomas-at-pnl.gov } } } } ---------- } From: Tom Phillips } Sent: Tuesday, April 10, 2001 11:22 AM } To: Alwyn Eades } Cc: Microscopy-at-sparc5.microscopy.com } Subject: Re: Scanners } } } ------------------------------------------------------------------------ } 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 too am about to buy and I would make a couple of comments on your } evaluation. First, let me remind everyone that the Dynamic range is } a log scale so small numerical differences are significant. } } I also think the Nikon Coolscan 8000 looks great but it only takes a } 2.5 x 3.5 negative which is smaller than my JEOL and Hitachi EM } negative sizes (~ 3 1/2 by 4 1/2"). Have these EM manufacturers gone } to a smaller film size or is Nikon using a non-Japanese EM as their } standard? seems odd but I don't see how the Nikon would be very } useful. You say a {2000 line scanner would be useful 9 out of 10 } times but want the 2000+ lines for the occasional high res scan. I } would argue that the size of the negative was the more important } variable to be worried about. The Nikon couldn't handle 4x5 LM } negatives or transparencies from autoradiography of } Westerns/Northerns, etc. } } My leading candidate is the ArtixScan 1100 has a Dmax of 3.9 (about } $1600 with SCSI card). This was has a 1000 x 2000 dpi resolution. } more details at www.microtek.com. This is my leading candidate. It } was 4 negative carriers and I await word whether one could be } modified to carry a 3 1/2 by 4 1/2 negative. At worst, I will have } my scientific instrumentation shop guys fabricate a holder. It comes } with a glass 8 x 10 glass carrier for odd size negs but I want to } avoid Newton rings and want a glassless carrier. } } I would appreciate comments on the following argument (I think I have } this correctly figured out but am not sure since so many out there } seem to want to have a higher resolution scanner). I have a Fuji } Pictrography 3000 printer with a 400 dpi output that is as good as } any other widely available printer in the academic world. If you } figure the maximum published image size is about 8 inches, that would } mean the maximum image size be 3200 dpi wide. A 1000 dpi scan of my } negative would be 4500 x 3500 dpi. I could crop by about 28% or 10% } depending on the orientation of the negative and still be taking full } advantage of the printer resolution. In reality, most EM publication } prints are smaller than 8" wide so one could crop even more and still } not need more than 1000 dpi. A resolution } 1000 dpi would be } useful for subtle morphometric analysis but a 4000 dpi scan of a 3 x } 4 negative would be 192 MB. That is pretty big for doing morphometry } on! A 1000 dpi scan of a 3.5 x 4.5" negative would be about 16 MB } and that is much more manageable. Perhaps the difference is in the } type of EM we are doing. I am working with biological specimens } doing standard thin section type stuff. are you doing some Material } Sci application that demands more? } } } I will be interested in Alwyn (and any others) reply since I hope to } buy one soon! } } } } . } } } } } } There was a thread recently on scanners for TEM film. I } have looked up } } all the models mentioned, on the web and called agents for } prices - and } } produced a comparative table, given below. } } } } I do not guarantee that the figures are accurate but they are my best } } interpretation of the data given. } } } } In the light of experience and Nestor's comments, I would suggest that } } 2000 dpi is a minimum for TEM negatives. You may be able to get away } } with less nine times out of ten, but there will be occasions when you } } need more. } } I would exclude the Minolta and all the Epsons from consideration } } (despite the incredibly low prices of some of the Epsons) because of } the } } low pixel density. } } } } Among the rest the Nikon has the best pixel density and the best } optical } } density (another critical parameter for TEM negatives). The price is } } very competitive too. The Nikon web site does not give a time for } } scanning a negative. On the face of it the Nikon would be a best buy } - } } get a separate, inexpensive flatbed scanner for the other work. } } } } These comments are all my own opinions based on manufacturers' data. } } Since we are considering purchase any comments to the } contrary would be } } most welcome. } } } } } } } } Code Maker Model Type } } } } } } } } A Agfa DuoScan T2500 Flatbed } } -Transparency included } } } } B Epson 1640 several versions Flatbed } } -Transparency option } } 1680 several versions } } } } C 1600 several versions Flatbed } } -Transparency included } } } } D Imacon Flextight Precision II Drum -for } } film and large format } } } } E Minolta Dimage ScanMulti II Film } } } } F Nikon Super Coolscan 8000ED Film } } } } G Polaroid 45 Ultra Film } } } } H Umax Powerlook 3000 Flatbed } } -Transparency included } } } } } } } } } } } } Code dpi OD Time Price } } Opinion } } at 6 x 9 cm } } } } } } A 2500 x2500 3.4 3 min } } $4,500 Fair } } } } B 1600 x 3200 3.6 } } $300-$3000 Poor } } } } $800-$1400 Poor } } } } C 1600 x 3200 3.3 } } $650-$1160 Not suitable } } } } D 2240 x2240** 3.9/4.1 N/A above } } $10k Good: low pixel density } } } } E 1128 x 1128 3.6 } } Not suitable } } } } F 4000 x 4000 4.2 N/A } } $2,695 V. Good } } } } G 2500 x 2500 3.8 5 min } } $7,495 Good but pricey } } } } H 3048 x 3048 3.6 3 min } } $6,499 Good } } } } } } -- } } .......... } } Alwyn Eades } } Department of Materials Science and Engineering } } Lehigh University } } 5 East Packer Avenue } } Bethlehem } } Pennsylvania 18015-3195 } } Phone 610 758 4231 } } Fax 610 758 4244 } } jae5-at-lehigh.edu } } -- } 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)
-- 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)
Does anyone know where I can find a multiple staining device for TEM grids using UA and Reynolds lead citrate that will result in CLEAN grids. Thank you Connie rosscac-at-okstate.edu
} Name: Kade } Organization: RMIT Melbourne Australia } Education: Graduate College } Location: Melbourne, Victoria, Australia } } Question: Hi, I have just purchased a microscope, a good biological } one. I need a microscope kit but nobody sells them around here. } Anyway already I have glass slides and covers. I have read some on } microscopy and I need an adhesive, resin I think its called to } prepare slides? is this true? } Also some ink to stain specimens. What are the names of all these } chemicals so I can buy them all seperatly since no one sells them all } together.
Kade -
You need a book on "microtechnique" plus a general idea of the types of specimens that you want to look at, BEFORE you start ordering things. Even tho you've "read some on microscopy " you could start with this book:
Nachtigall, W. 1995 Exploring With the Microscope 160 pp. hardbound. 6.5x9.5". $19.95. ISBN 0-8069-0866-1 Sterling Publishing Co., NY. Although this book is intended for adult amateur microscopists, it is well written and will provide teachers and classroom volunteers with much useful information on "serious" light microscopy. Almost half of the book is devoted to simple preparation methods for biological specimens and descriptions (with gool illustrations) of commonly encountered organisms. Adult. RECOMMENDED
You will find a lot of help on the amateur microscopy website http://www.microscopy-uk.org.uk
Both of these listings are from the Project MICRO bibliography (URL below).
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
Your information is correct and mine is not. The Dmax of the 8700 is 3.4.
Larry
---------- From: Tom Phillips Sent: Wednesday, April 11, 2001 8:22 AM To: Microscopy-at-sparc5.microscopy.com Cc: jae5-at-lehigh.edu; Thomas, Larry (PNNL) Subject: RE: Scanners
I would love to take advantage of the Firewire option but my information is that the 8700 has a Dmax of 3.4 vs the 3.9 for the 1100. That is a significant difference. Do EM negatives of biological thin sections reach that? I think so. I do a lot of EM immunocytochemistry and have to look for gold (intensely black) against a very dark tissue component so I am hoping the higher Dmax improves my results. I frequently scan negatives on a Umax 1100 (Dmax 3.4??) and can't differentiate the gold from the background although by eye I can discriminate them when the negative is placed on a light box. Changing my exposure would give me an unuseable image for the rest of the tissue. Maybe this is an extreme case but I suspect that lots of "dark organelles" (e.g., lysosomes, nuclei) have fine structure that get lost in the scanning with a low Dmax scanner. Tom
} A colleague and I each recently bought Microtek scanners to scan TEM } negatives. } I have the Artixscan 1100 and he has the Model 8700 which has similar } characteristics (actually higher resolution -1200dpi), 3.9 dmax at } 42 bits color } (14 grayscale), and the glassless film carrier setup. The 8700 has USB and } Firewire interfaces and is cheaper ( {$1000), and the 1000 dpi Model 1100 has a } SCSI interface. You might want to check out the specs of the lower cost model } 8700 on the microtekusa website if your computer can handle USB or Firewire. } Both scanners have performed up to our expectations, which I would } characterize } as modest. Microtek does not supply a 3-1/4 x 4 " negative carrier } for standard } size TEM film but you can easily make a serviceable one from stiff paper or } light cardboard. } } How much scanner resolution should you buy? The answer depends on how you } intend to use it. Most applications do not require capturing the full } resolution of the negative. From a practical viewpoint, the scanner } resolution } just determines how many times you can magnify the negative image to } produce the } final print size. For example, to get a publication-size print at 300 dpi, an } image scanned at 1200 dpi scan could be zoomed 4X. A practical alternative to } spending more for higher scanning resolution is to take photos at higher } magnification. One exception is with lattice images from the TEM, which } (depending on the lattice fringe spacing on the negative) might require higher } scan resolutions to avoid getting a moire effect. (Of course, not everyone } agrees. My colleague prefers to always scan at the maximum resolution). } } What does a Dmax of 3.9 mean to you? To me it means a very dark } negative. D is } the log of the transmitted to incident intensity ratio. I wonder if } users ever } actually verify the manufacturer's specs with a calibrated density target. A } Dmax of 3.9 can be useful for scanning TEM diffraction patterns that } might have } high contrast, but TEM micrograph negatives of metals and ceramics generally } don't have that much contrast and biological thin section photos tend to have } rather weak contrast. If your negatives are simply dark, use shorter photo } exposure times. Scanning with maximum allowed grayscale resolutions (e.g., 14 } bits rather than 8) is highly recommended if you intend to enhance or adjust } images, but that's another story. } } } Larry Thomas } Pacific Northwest National Laboratory } MSIN P8-16 } P.O. Box 999 } Richland, WA 99352 } Phone: (509)372-0793 Fax: (509)376-6308 } Email: mailto:Larry.Thomas-at-pnl.gov } } } } ---------- } From: Tom Phillips } Sent: Tuesday, April 10, 2001 11:22 AM } To: Alwyn Eades } Cc: Microscopy-at-sparc5.microscopy.com } Subject: Re: Scanners } } } ------------------------------------------------------------------------ } 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 too am about to buy and I would make a couple of comments on your } evaluation. First, let me remind everyone that the Dynamic range is } a log scale so small numerical differences are significant. } } I also think the Nikon Coolscan 8000 looks great but it only takes a } 2.5 x 3.5 negative which is smaller than my JEOL and Hitachi EM } negative sizes (~ 3 1/2 by 4 1/2"). Have these EM manufacturers gone } to a smaller film size or is Nikon using a non-Japanese EM as their } standard? seems odd but I don't see how the Nikon would be very } useful. You say a {2000 line scanner would be useful 9 out of 10 } times but want the 2000+ lines for the occasional high res scan. I } would argue that the size of the negative was the more important } variable to be worried about. The Nikon couldn't handle 4x5 LM } negatives or transparencies from autoradiography of } Westerns/Northerns, etc. } } My leading candidate is the ArtixScan 1100 has a Dmax of 3.9 (about } $1600 with SCSI card). This was has a 1000 x 2000 dpi resolution. } more details at www.microtek.com. This is my leading candidate. It } was 4 negative carriers and I await word whether one could be } modified to carry a 3 1/2 by 4 1/2 negative. At worst, I will have } my scientific instrumentation shop guys fabricate a holder. It comes } with a glass 8 x 10 glass carrier for odd size negs but I want to } avoid Newton rings and want a glassless carrier. } } I would appreciate comments on the following argument (I think I have } this correctly figured out but am not sure since so many out there } seem to want to have a higher resolution scanner). I have a Fuji } Pictrography 3000 printer with a 400 dpi output that is as good as } any other widely available printer in the academic world. If you } figure the maximum published image size is about 8 inches, that would } mean the maximum image size be 3200 dpi wide. A 1000 dpi scan of my } negative would be 4500 x 3500 dpi. I could crop by about 28% or 10% } depending on the orientation of the negative and still be taking full } advantage of the printer resolution. In reality, most EM publication } prints are smaller than 8" wide so one could crop even more and still } not need more than 1000 dpi. A resolution } 1000 dpi would be } useful for subtle morphometric analysis but a 4000 dpi scan of a 3 x } 4 negative would be 192 MB. That is pretty big for doing morphometry } on! A 1000 dpi scan of a 3.5 x 4.5" negative would be about 16 MB } and that is much more manageable. Perhaps the difference is in the } type of EM we are doing. I am working with biological specimens } doing standard thin section type stuff. are you doing some Material } Sci application that demands more? } } } I will be interested in Alwyn (and any others) reply since I hope to } buy one soon! } } } } . } } } } } } There was a thread recently on scanners for TEM film. I } have looked up } } all the models mentioned, on the web and called agents for } prices - and } } produced a comparative table, given below. } } } } I do not guarantee that the figures are accurate but they are my best } } interpretation of the data given. } } } } In the light of experience and Nestor's comments, I would suggest that } } 2000 dpi is a minimum for TEM negatives. You may be able to get away } } with less nine times out of ten, but there will be occasions when you } } need more. } } I would exclude the Minolta and all the Epsons from consideration } } (despite the incredibly low prices of some of the Epsons) because of } the } } low pixel density. } } } } Among the rest the Nikon has the best pixel density and the best } optical } } density (another critical parameter for TEM negatives). The price is } } very competitive too. The Nikon web site does not give a time for } } scanning a negative. On the face of it the Nikon would be a best buy } - } } get a separate, inexpensive flatbed scanner for the other work. } } } } These comments are all my own opinions based on manufacturers' data. } } Since we are considering purchase any comments to the } contrary would be } } most welcome. } } } } } } } } Code Maker Model Type } } } } } } } } A Agfa DuoScan T2500 Flatbed } } -Transparency included } } } } B Epson 1640 several versions Flatbed } } -Transparency option } } 1680 several versions } } } } C 1600 several versions Flatbed } } -Transparency included } } } } D Imacon Flextight Precision II Drum -for } } film and large format } } } } E Minolta Dimage ScanMulti II Film } } } } F Nikon Super Coolscan 8000ED Film } } } } G Polaroid 45 Ultra Film } } } } H Umax Powerlook 3000 Flatbed } } -Transparency included } } } } } } } } } } } } Code dpi OD Time Price } } Opinion } } at 6 x 9 cm } } } } } } A 2500 x2500 3.4 3 min } } $4,500 Fair } } } } B 1600 x 3200 3.6 } } $300-$3000 Poor } } } } $800-$1400 Poor } } } } C 1600 x 3200 3.3 } } $650-$1160 Not suitable } } } } D 2240 x2240** 3.9/4.1 N/A above } } $10k Good: low pixel density } } } } E 1128 x 1128 3.6 } } Not suitable } } } } F 4000 x 4000 4.2 N/A } } $2,695 V. Good } } } } G 2500 x 2500 3.8 5 min } } $7,495 Good but pricey } } } } H 3048 x 3048 3.6 3 min } } $6,499 Good } } } } } } -- } } .......... } } Alwyn Eades } } Department of Materials Science and Engineering } } Lehigh University } } 5 East Packer Avenue } } Bethlehem } } Pennsylvania 18015-3195 } } Phone 610 758 4231 } } Fax 610 758 4244 } } jae5-at-lehigh.edu } } -- } 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)
-- 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)
Dear all, We are having a great deal of difficulty getting hold of our normal film "Tritiated Hyper film" for use with tritiated ligands. Does any one out there know of a local supplier (U.K.)
We use 70 % isopropanol to clean emersion oil off of our lenses.
Dotty Sorenson Microscopy and Image Analysis Laboratory Department of Cell and Developmental Biology University of Michigan Medical School Ann Arbor, Michigan (734)763-1170 FAX (734)763-1166 dsoren-at-umich.edu
On Wed, 11 Apr 2001 mckaylodge-at-aol.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 } -----------------------------------------------------------------------. } } } } Email: mckaylodge-at-aol.com } Name: Robert Lodge } } Organization: McKay Lodge Home School } } Education: 9-12th Grade High School } } Location: Oberlin, OH 44074 } } Question: My student accidently got immersion oil on the 40x Leica } Plan Acromat. I took a chance and used a fine artist's brush and } xylene to clean it recalling (I may be wrong) that lens adhesives are } soluble in alcohols not xylene, toluene and similar. Well, the lens } didn't fall out. Too bad because I need an excuse to upgrade! If (or } when) this happens again, what would you recommend for cleaning? } } Bob Lodge } } --------------------------------------------------------------------------- }
Has anyone had experience in determining the concentration of particles in a solution by counting on EM grids in a similar way to using a haemocytometer? Is it possible to count the particles in a defined number of grid squares then calculate back to the area of the grid and the amount of solution which was applied and allowed to dry down?
Richard Gardiner Department of Plant Sciences University of Western Ontario
Just tried to order some Kodabrome II RC paper, grade F5. Vendor says it is discontinued. Any suggestions for an alternative? We have been using Kodabrome RC II for a long time with a Mohr processor. How about Polycontrast RC? Or is this the beginning of the end for old fashioned darkrooms?
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
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One of the big advantages of computer printing of TEM images is the ability to adjust the contrast of the final image. However, to prevent generating a negative that has completely overexposed areas which are difficult to salvage by any means-such as one may get with metal or ceramic specimens-an alternative film developer may help. There are two bath "split" developers which lower contrast to manageable levels, in effect chemically "dodging" a developing negative. Fixing is carried out normally. I'm told biological specimens don't usually need such treatment. I have no financial interest in the following company, but I am pleased with the results obtained with their developer called Diafine. It is made by Acufine Inc., 5441 North Kedzie Ave. Chicago, Il., 60625.
Bernard Kestel E-mail: {kestel-at-anl.gov} Materials Science Division Argonne National Laboratory Argonne, Il., 60439
by ultra5.microscopy.com (8.9.3+Sun/8.9.1) id TAA26206 for dist-Microscopy; Wed, 11 Apr 2001 19:12:49 -0500 (CDT) Received: from no_more_spam.com (sparc5 [206.69.208.10]) by ultra5.microscopy.com (8.9.3+Sun/8.9.1) with SMTP id TAA26197 for "MicroscopyFilteredEmail1-at-msa.microscopy.com"; Wed, 11 Apr 2001 19:12:19 -0500 (CDT) Received: from [206.69.208.21] (mac21.zaluzec.com [206.69.208.21]) by ultra5.microscopy.com (8.9.3+Sun/8.9.1) with ESMTP id TAA26190 for {Microscopy-at-MSA.Microscopy.Com} ; Wed, 11 Apr 2001 19:12:08 -0500 (CDT) Mime-Version: 1.0 X-Sender: zaluzec-at-ultra5.microscopy.com Message-Id: {p05001900b6faa35d2003-at-[206.69.208.21]}
Below is the result of your feedback form. It was submitted by (jgh7f0-at-mizzou.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Wednesday, April 11, 2001 at 14:32:21 ---------------------------------------------------------------------------
Email: jgh7f0-at-mizzou.edu Name: john harris
Organization: university of missouri-columbia
Education: Undergraduate College
Location: Columbia, Missouri
Question: Dear Sir or Madam; i am looking for an image of S. Agalactiae attached to an epithelial cell or some other gram + cocci.
Kodabrome is for sure an old product. But a good one. I quit using it long ago in favor of Ilford paper. Ilford makes RC paper--but my work has only been output to archival fiber paper. But I suspect that the RC quality is still up to par.
I use plain matte for normal prints--and archival matte fiber for fine art prints (nudes, etc.). I use a Kreonite processor for the print papers. All b/w neg media is hand processed in separate roll holders (Nikor). My normal format is 6x4.5cm and 6x7cm. The same recipes would apply to other formats.
I did some 4x5" work in prior years and do the same mechanics today.
Try some Ilford paper.
gary g.
http://photoweb.net
At 03:25 PM 4/11/2001, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Wow...this is really deja vu. I used Diafine and Acufine way many years ago. At that time, the issue was push processing. Today, the zone system prevails. Using the zone system does not require special chemicals. It is a matter of how the neg is rated and how it is processed.
Look up some of Ansel Adams' works (The Negative). The main idea is to adjust your neg's EV for total tonal range such that it can be printed with less than bone crushing effort.
All of my fine art prints are shot and printed using this process.
gary g.
http://photoweb.net
At 04:41 PM 4/11/2001, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I used Illiford RC paper for a long time and I liked it better than anything else I have ever used. It may be just be personal preference but it seemed to have brighter higlights and blacker blacks than I could get with other RC papers. Just be sure and get Illford filters to go with it. One other nice thing is it has a matt finsh as well as glossy if you don't want glossy prints. The matt will scan great. It doesn't have texture problems like most pearl or simi-glossy papers have. The matt finish displays a lot better than glossy finsh IMHO.
Gordon Gordon Couger gcouger-at-couger.com Stillwater, OK www.couger.com/gcouger
} From: "Gary Gaugler" {gary-at-gaugler.com}
} Kodabrome is for sure an old product. But a good one. } I quit using it long ago in favor of Ilford paper. Ilford } makes RC paper--but my work has only been output to archival } fiber paper. But I suspect that the RC quality is still up to par. } } I use plain matte for normal prints--and archival matte fiber } for fine art prints (nudes, etc.). I use a Kreonite processor } for the print papers. All b/w neg media is hand processed } in separate roll holders (Nikor). My normal format is 6x4.5cm } and 6x7cm. The same recipes would apply to other formats. } } I did some 4x5" work in prior years and do the same } mechanics today. } } Try some Ilford paper. } } gary g. } } http://photoweb.net } } } } } Just tried to order some Kodabrome II RC paper, grade F5. Vendor says it is } } discontinued. Any suggestions for an alternative? We have been using } } Kodabrome RC II for a long time with a Mohr processor. How about } } Polycontrast RC? Or is this the beginning of the end for old fashioned } } darkrooms? } } } } Jonathan Krupp
} I am using Polymax II RC. For draft pictures it's good enough in my point } of view. In compare with Ilford Multigrade III RC, Polymax gives better } contrast (filter #5 on Ilford is equal to #3 on Polymax in my hands).
Sergey
} At 03:25 PM 4/11/2001, you wrote: } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
_____________________________________
Sergey Ryazantsev Ph. D. Electron Microscopy UCLA School of Medicine Department of Biological Chemistry Box 951737 Los Angeles, CA 90095-1737
Usualy to reduce contrast you under expose the film. Then develop it to the disired density using a developer that generates low contrast. One way to reduce contrast is to dilute you developer by a factor of 2, 4 or more with water. It extends the developing time a good deal but it reduces the contrast. You might also look at low contrast developers that work with the film you are using.
A few question on rec.photo.darkroom will get you more information than you can handle and some of it will actually work.
Gordon Gordon Couger gcouger-at-couger.com Stillwater, OK www.couger.com/gcouger
} } } } } } One of the big advantages of computer printing of TEM images is the } } ability to adjust the contrast of the final image. However, to prevent } } generating a negative that has completely overexposed areas which are } } difficult to salvage by any means-such as one may get with metal or ceramic } } specimens-an alternative film developer may help. } } There are two bath "split" developers which lower contrast to } } manageable levels, in effect chemically "dodging" a developing negative. } } Fixing is } } carried out normally. I'm told biological specimens don't usually need } } such treatment. } } I have no financial interest in the following company, but I am } } pleased with the results obtained with their developer called Diafine. It is } } made by Acufine Inc., 5441 North Kedzie Ave. Chicago, Il., 60625. } } } } Bernard Kestel E-mail: {kestel-at-anl.gov} } } Materials Science Division } } Argonne National Laboratory } } Argonne, Il., 60439 } }
-----Original Message----- } From: christine [mailto:ac.richardson2-at-btinternet.com] Sent: Wednesday, April 11, 2001 1:59 PM To: microscopy-at-sparc5.microscopy.com
Dear all, We are having a great deal of difficulty getting hold of our normal film "Tritiated Hyper film" for use with tritiated ligands. Does any one out there know of a local supplier (U.K.)
Back in the pre PC (chemical) days, toluene was a recommended solvent for cleaning lenses of immersion oil. Now, we at National Steel, wipe off excess oil with lens tissue and then clean the lens with Kodak lens cleaner solution.
Best regards,
Sam Purdy National Steel Tech Center Trenton MI
} ---------- } From: mckaylodge-at-aol.com } Sent: Wednesday, April 11, 2001 9:59 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: Ask-A-Microscopist:Help Cleaning Lenses } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } } Email: mckaylodge-at-aol.com } Name: Robert Lodge } } Organization: McKay Lodge Home School } } Education: 9-12th Grade High School } } Location: Oberlin, OH 44074 } } Question: My student accidently got immersion oil on the 40x Leica } Plan Acromat. I took a chance and used a fine artist's brush and } xylene to clean it recalling (I may be wrong) that lens adhesives are } soluble in alcohols not xylene, toluene and similar. Well, the lens } didn't fall out. Too bad because I need an excuse to upgrade! If (or } when) this happens again, what would you recommend for cleaning? } } Bob Lodge } } -------------------------------------------------------------------------- } - }
Kodabrome has not been discontinued across the board. However, certain size/surface/grade combinations have been. Kodabrome II RC F5 is available in 250sheet 8x10 packages only. We stock them but a dealer should be able to order them. As far as alternatives.... Agfa offers Brovira Speed RC, a developer incorporated paper in grades 2-5 and available in 100 sheet 8x10 packs. Brovira Speed is a cold tone paper. Agfa also offers a Multicontrast RC product. From Kodak, Polycontrast or Polymax are variable contrast papers. Polycontrast is developer incorporated, as Kodabrome is, so it will process in developers as well as many activators. Polymax requires the use of a developer and will not process in activators. Ilford offers Multigrade IV Deluxe which is not developer incorporated. All of these papers use filters to control contrast, although a #5 filter with any of them is not the same as a grade 5 Kodabrome. Both Polymax and Multigrade IV have a slightly wider contrast range than Polycontrast. For those who prefer a cooler tone print, Ilford also has a new paper, Multigrade Cooltone, a non developer incorporated paper with a cooler image tone and a cool white base tint.
George
George Laing National Graphic Supply v:(800) 223-7130 X3109 f:(800) 832-2205 email: scisales-at-ngscorp.com
} Just tried to order some Kodabrome II RC paper, grade F5. Vendor } says it is discontinued. Any suggestions for an alternative? We have } been using Kodabrome RC II for a long time with a Mohr processor. How } about Polycontrast RC? Or is this the beginning of the end for old } fashioned darkrooms? } } Jonathan Krupp
The paper is still listed on the Kodak Web sight, http://www.kodak.com/cluster/global/en/professional/support/techPubs/f33/old/f33.shtml . The web sight does list Kodabromide paper as discontinued but lists Kodabrome II RC as its replacement and does not list the latter as discontinued. I was curious, so, I called the 800 technical hotline (800) 242-2424 (option 02 gives you an operator) and asked. They said that it is still available but not in as many sizes as it use to be.
I've had venders tell me a product is discontinued when it is only discontinued from their stocking process, not by the manufacture. In this case it may be that only the size you wanted was discontinued. You may want to call the vendor back or check with another vender.
I have no connection with Kodak but wanted to see if the predictions I'd heard about, as you put it, " the beginning of the end for old fashioned darkrooms," was starting. It would seem not quite yet in this case. Though I'm hearing that in 2 or 3 years it will, somewhat sad I think.
Jim Roberts
James L. Roberts Firearm and Toolmark Examiner Ventura Co. Sheriff's Lab (805) 654-2308 James.Roberts-at-mail.co.ventura.ca.us
} } } {jmkrupp-at-cats.ucsc.edu} 04/11/01 03:25PM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi:
Just tried to order some Kodabrome II RC paper, grade F5. Vendor says it is discontinued. Any suggestions for an alternative? We have been using Kodabrome RC II for a long time with a Mohr processor. How about Polycontrast RC? Or is this the beginning of the end for old fashioned darkrooms?
Jonathan Krupp Microscopy & Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 jmkrupp-at-cats.ucsc.edu
Yes, Thermanox cover slips survive treatment with HMDS. We occationally process cell monlayers grown on them for SEM.
Regards,
Dotty
Dotty Sorenson Microscopy and Image Analysis Laboratory Department of Cell and Developmental Biology University of Michigan Medical School Ann Arbor, Michigan (734)763-1170 FAX (734)763-1166 dsoren-at-umich.edu
On Thu, 12 Apr 2001, Randall, Kevin J 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 need to process some thermanox coverslips for SEM. Does anyone know } whether thermanox survives HMDS? } } Cheers } } Kevin }
Hi All: Thank you all for sharing your experiences about working with SiC. Your suggestions have helped me come up with a plan of attack. Regards, Mike Coviello UT Arlington
We are a Kodak distributor and we checked with Kodak this morning and Kodabrome II RC paper, grade F5 is still available. If you still interested you may contact me off-line.
John Arnott
Jon Krupp 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: } } Just tried to order some Kodabrome II RC paper, grade F5. Vendor says it is } discontinued. Any suggestions for an alternative? We have been using } Kodabrome RC II for a long time with a Mohr processor. How about } Polycontrast RC? Or is this the beginning of the end for old fashioned } darkrooms? } } Jonathan Krupp } Microscopy & Imaging Lab } University of California } Santa Cruz, CA 95064 } (831) 459-2477 } jmkrupp-at-cats.ucsc.edu
We are about to install a Kimball ES-423E (extended life) LaB6 cathode in a Hitachi H7100 TEM and were wondering if anyone had some starting points for the voltage settings for filament heating. This would save us a lot of time, if so.
Thank you.
John B.
-- ############################################################## John J. Bozzola, Ph.D., Director I.M.A.G.E. (Integrated Microscopy & Graphics Expertise) 750 Communications Drive - MC 4402 Southern Illinois University Carbondale, IL 62901 U.S.A. Phone: 618-453-3730 Fax: 618-453-2665 Email: bozzola-at-siu.edu Web: http://www.siu.edu/departments/shops/cem.html ##############################################################
Here is an odd way that I thought up to clean oil from an objective without using solvents that may attack lens cement.
Remove the lens from the scope. I usually like to view the oil contamination using a stereoscopic microscope. Wipe excess oil away - I like using Ross Optical Paper. Using a cotton applicator wrapped in Ross optical paper, apply a small amount of Dawn dishwashing detergent. Gently work the surface to emulsify the oil into the detergent using the optical papered applicator. You may have to add a small amount of water. Do not allow fluids to go much beyond the lens area!
At this point, hold the lens vertical with the back focal plane pointing up. Apply a small amount of deionized water on the final lens element from the side of the opjective to create a hanging drop. I usually use a wash bottle or 10cc syringe. The surface tension of water will create a small drop around the optical surface. Start a stream of DI water flowing through the small drop to wash away the oil/water suspension. After about a minute, stop the water flow while allowing a small drop of water to stay on lens. At this time, blow the water off using C02 or some other clean compressed gas. This will eliminate streaks. Upon inspection, if you see contamination, repeat the process and your problem should be solved.
It's weird but it works.
Robert
Robert Fitton Teaching Associate/Director of Laboratories Luther College Department of Biology 700 College Drive Decorah, IA 52101
Voice 563-387-1559 FAX 563-387-1080
Enjoy a visit to our website: http://www.luther.edu/~biodept/
Dear Richard, The most common use of this is for asbestos fibre load determination. I did it once. You count the number of fibres in a specified number of grids openings, then calculate back to the original collected volume or vacuumed area. It should work similarly for any recognizable particle. I could look up the method, if you like. At 06:08 PM 4/11/01 -0400, you wrote: } } Has anyone had experience in determining the concentration of particles } in a solution by counting on EM grids in a similar way to using a } haemocytometer? Is it possible to count the particles in a defined } number of grid squares then calculate back to the area of the grid and } the amount of solution which was applied and allowed to dry down? } } Richard Gardiner
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
Below is the result of your feedback form. It was submitted by (dngeorge-at-wam.umd.edu) from http://www.msa.microscopy.com/Ask-A-Microscopist.html on Thursday, April 12, 2001 at 16:04:35 ---------------------------------------------------------------------------
Email: dngeorge-at-wam.umd.edu Name: Damali Martin
Organization: University of Maryland
Education: Graduate College
Location: College Park, MD
Question: I have embedded and sectioned some salivary glands in epon and have placed the sections on glass slides. However, when I counterstain, a high percentage of sections float off and are lost. How can this problem be prevented?
Also, several of my sections have wrinkles in them. Is there any trick to getting rid of them?
Anyone got any suggestions as to the cause of (and remedy for) the pronounced image shift that occurs when I turn the Y stigmator control on my JEOL 840A?
It also has the usual stigmatic effect.
The X control shifts the image only very little.
The coils seem to check out OK.
There's something quite poetic about working on this particular part of the instrument today of all days.
thanks
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
Stigmators operate as two opposing coils. It may be that one of the coils for the Y stigmator is not active while the opposing Y coil is. Perhaps one of the coils has shorted? The resistance of these coils is very low, making a simple check rather difficult, not to mention that they are often series connected within the column making physical access difficult. But if you can measure the current drawn by each, you may find a significant difference if one is shorted.
If so, the only remedy is replacement of the coils.
On Friday, April 13, 2001 1:12 PM, Ritchie Sims [SMTP:r.sims-at-auckland.ac.nz] wrote: } } Hi } } Anyone got any suggestions as to the cause of (and remedy for) the } pronounced image shift that occurs when I turn the Y stigmator } control on my JEOL 840A? } } It also has the usual stigmatic effect. } } The X control shifts the image only very little. } } The coils seem to check out OK. } } There's something quite poetic about working on this particular part } of the instrument today of all days. } } thanks } } rtch } } } Ritchie Sims Phone : 64 9 3737599 ext 7713 } Department of Geology Fax : 64 9 3737435 } The University of Auckland email : r.sims-at-auckland.ac.nz } Private Bag 92019 } Auckland } New Zealand } }
Allen R. Sampson, Owner Advanced Research Systems 317 North 4th. Street St. Charles, Illinois 60174 voice 630.513.7093 fax 630.513.7092
About 25 years ago, when I was taking the embryology course at the Marine Biology Laboratory at Woods Hole, we were taught the following methods for cleaning oil from lenses by Robert Allen, a well known microscopist:
Remove the lens from the microscope and invert it. Remove most of the excess oil from the objective with lens paper without touching the objective surface itself. Gently lay a strip of lens paper over the objective. Since the lens is recessed the lens paper does not touch the glass directly. Then place a drop or two of ether on the lens paper just next to where it contacts the lens, and draw the lens paper over the objective surface. The ether vapors swirl around under the lens paper and dissolve the oil while the lens paper absorbs the mixture and carries it away, without actually touching the glass surface. You might need to repeat a few times to remove the oil.
It works with all but the most stubborn deposits. The upside is that it completely avoids touching the glass surface, and the ether fumes are in contact with the element such a short time that it miakes it less likely you will dissolve the glues that hold the lens elements together. The downside is that it requires ether. I haven't tried it with other solvents. -- Gary P. Radice gradice-at-richmond.edu Associate Professor of Biology 804 289 8107 (voice) University of Richmond 804 289 8233 (FAX) Richmond VA 23173 http://www.science.richmond.edu/~radice
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Accountabilities % of time
70 Collect data from production activities to track and improve quality. Develop collection systems to optimize this process. We have a new Hitachi S3000N with an Oxford INCA EDS system and an Oxford/Gatan CL system with cryostage. Routine analysis will include EBIC, EDS, and CL.
20 Interface with Manufacturing Engineers, Application engineers, Detail Engineers, Product Managers, Technicians, Maintenance, Tool Room, Field Service, and other stakeholders to achieve assigned project goals. Recommend design changes to improve manufacturability and quality.
10 Coordinate project work activity, including testing, machining, and purchasing, as needed to ensure meeting project schedules. Design and build fixtures, prototypes, and samples. Provide training on equipment and techniques to assemblers.
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Learn about ADC - The Broadband Company at www.adc.com
I usually see image shift as a result of stigmation in both of our scopes (JEOL 840A and Hitachi 2460N). I rather accepted that it just went with the territory. If it is a sign of a problem with the scope or its alignment, I too would be interested in hearing how to eliminate it.
Warren
At 06:12 PM 4/13/2001 +0000, you wrote:
} Hi } } Anyone got any suggestions as to the cause of (and remedy for) the } pronounced image shift that occurs when I turn the Y stigmator } control on my JEOL 840A? } } It also has the usual stigmatic effect. } } The X control shifts the image only very little. } } The coils seem to check out OK. } } There's something quite poetic about working on this particular part } of the instrument today of all days. } } thanks } } rtch } } Ritchie Sims Phone : 64 9 3737599 ext 7713
---------------------- Warren E. Straszheim Materials Analysis and Research Lab Iowa State University 23 Town Engineering Ames IA, 50011-3232
The 840 has some pots (potentiometers) on one of the circuit boards that adjust out the image shift while adjusting the stigmators. They probably balance a bias, or an offset, in the circuit for the stigmation coils. If you have the manual with the schematics, they may be identified. I may be able to find it in ours, if you don't have it.
Normally, X or Y stig adjustment will cause image shifting. More shifting at higher mag. Since stigmation is done with coils by the scan coils, changing current in the stig coils appears to the beam as a change in scan coil current. Hence, the image shifts. Later day systems account for this by feeding some of the stig signal to the scan coils as feedback. It will also accept additional feedback from magnification. The operation is to adjust scan coil current and condenser current as stig is adjusted so that the net result is minimal image shift with varying stig input.
Since one stig channel out of two is not working right, there are likely one of two or three things which could go wrong. First, and worst, the stig coil is open or shorted. Somewhat unlikely I think. Second, one part of the stig coil drive circuit has failed. Third, some part of the feedback system has failed. Since you say that it does stig, but causes image shift, then the stig circuit itself and the coil is OK. You can verify this.
Since the stig coil drive systems are identical (ususally), you should be able to trace readings from the good channel and compare them to the bad channel. This should show right away where the problem is. The stig, beam alignment and image shift circuits are usually all the same. If so in your system, they will give plenty of data points for checking. The coils (one for X, one for Y) are typically driven by push-pull power transistors which are high current buffers on the output of a small op amp. The coils are in the negative feedback loop of the op amp. One lead of a coil would connect to the output of the buffer transistors (large ones) while the other end connects to the inverting input of the op amp and is returned to ground through a low value resistor. Measuring the voltage across this resistor will tell how much current is flowing through the coil (I=E/R).
Stig effect is lower at lower mags. So there would be less automatic compensation for stig vs. mag. Try a low mag setting and measure the voltage on each side of the stig coils (two leads each) and the voltage on the low value resistors. Have the stig controls at 12 O'clock each. If these readings match, odds are that the coils are for sure OK. Then the problem ought to be narrowed to the stig balance circuit. Look for this circuit and compare the two stig signal feedback loops for differences. It could be something as simple as a bad op amp in the path from the stig circuit to the beam alignment coil drivers. Since stig has little effect at low mag, but huge effect at high mag, the usual control path for stig compensaton versus magnification is to electronically change the beam position by sending a stig sourced voltage to the scan coil driver circuit.
If you don't have schematics for the system, that is of course a major problem. In this case, perhaps someone who has your same model has encountered this problem before and knows of the failure mechanism and cause.
gary g.
At 11:12 AM 4/13/2001, you wrote:
} Hi } } Anyone got any suggestions as to the cause of (and remedy for) the } pronounced image shift that occurs when I turn the Y stigmator } control on my JEOL 840A? } } It also has the usual stigmatic effect. } } The X control shifts the image only very little. } } The coils seem to check out OK. } } There's something quite poetic about working on this particular part } of the instrument today of all days. } } thanks } } rtch } } } Ritchie Sims Phone : 64 9 3737599 ext 7713 } Department of Geology Fax : 64 9 3737435 } The University of Auckland email : r.sims-at-auckland.ac.nz } Private Bag 92019 } Auckland } New Zealand
Stigmator shift is almost always down to the balance of the stigmator coils. Stigmators have 8 coils in four sets of two. If an opposing pair are not correctly balanced the stronger coil will cause the image to move away from that coil.
Some instruments have the balancing potentiometers easily accessible others hide them away in the electronics.
The pots will be called Xx, Xy, and Yx, Yy I do not know where they are on a JEOL 840 but if you look inside you may be lucky?
To adjust - 1. Place an easily recognizable feature in the center of the screen 2. Turn the X stigmator fully in one direction and re center with the Xx or Xy control whichever fits. 3. Turn in the opposite direction and use the other compensation control to center 4. Repeat with the Y stigmator and Yx and Yy controls.
I have just completed the promised "Monitoring & Maintaining Electron Microscope Performance" interactive CD, this area is covered in the instrument tuning section so its pretty fresh in my mind. Need to see even more for yourself then we have the course of the same name running in Sydney early October this year?
Kindest regards and good luck
Steve Chapman Senior Consultant, Protrain For professional training in SEM, TEM and EDX world wide www.emcourses.com
I have a Hitachi SEM model S-570 (1985). I am interested of getting a X-ray analyzer for this microscope. My preference would be to buy a used analyzer with an ultrathin window for light elements analysis.
If you know someone who is interested in selling it's analyzer, please let me know...
Thank you
Christian Normand Tekna Systems Plasma (819) 820-2204
I hesitated to make any response regarding this subject. But--here goes anyway. Someone may find it useful.
I think that the issue is not reduction of contrast but rather extension of tonal range of the negative. That is what the zone system is all about. I think that it is a fact that if the silver is not exposed, there is not going to be any information gleaned from that unexposed or too underexposed area. The approach I use for fine art images is to overexpose and under-develop. It takes a LOT of work to develop a complete system of ISO rating, development and printing to achieve marketable prints. But it certainly can be done. The goal is to minimize the time spent in the darkroom doing printing. Ideally, the neg should print without any effort on grade 3 paper. I only use Ilford archival matte fiber paper, so extensions to RC papers may or may not directly apply. I also do not use variable contrast paper. But the principles are extensible I would think.
Being all-digital with my SEM, I use realtime histogram feedback to adjust gain and dynamic range. For TEM, I have no direct experience. But I might suggest that the same photo techniques for fine art neg work may apply to TEM negs. If you think that this might be true, read on. Otherwise, nevermind.
A neg with bad dynamic range (huge extremes of EV or low contrast) is not going to produce a very good scan. This is the realm of drum scanners with 4.0-6.0D. Pixel resolution is subordinate to D range in this case. If one is going to print a neg on paper, that is one aspect of the problem. If the neg is to be output to a magazine or printed page, that is another aspect. So the crux of the matter is what the actual intended end use of the neg is to be? If it is a nice print, OK. If it is a magazine or litho output, these are 133lpi. Doing scans at 4000dpi only to print them in a magazine at 133lpi is rather silly. One would probably be better off just printing the neg and scanning at 300dpi.
But the 300 dpi is the effective dpi for the size of the final image....not the original neg. So, if the neg is say 3" x 4" and is to be printed at that same size, a 300 dpi scanner should do the job (ignoring tonal range for the time being). If the neg is to be printed at twice its physical size, then the scanner has to have twice the resolution (600 dpi). And it goes on from there. Thus, for most image output methods on a printed page, 600-1200 optical dpi should be plenty of resolution. If you agree with this discussion at this point, then the resolution factor should no longer be an issue. What remains is D range.
(Note that commercial images are created at high resolution to accommodate manipulation and merging with other images. This way, image quality can gradually be reduced without affecting the final result.)
But the negative is the key item in all of this discussion. A bad neg is not likely to do anyone much good. So the challenge is to get a good neg at the get go. Here we go, back to the zone system. The idea is to use the neg as a non-linear image capture media for data which may span a wide dynamic range. And in so doing, be able to output (print or scan) this information with fidelity and minimal effort. This drives D value.
An unexposed, developed neg will have a transmission of 100%. It is actually slightly lower due to absorption by the medium, emulsion and residual anti-dispersion coating. But at 100% transmission, the neg would have an opacity of 100%input/100%output or 1.0. Since D=log1/T, the D value for the unexposed neg is log 1/1=0.0. This then is Dmin. The area with the most exposure (darkest region) will pass the least amount of transmitted light. If, for example, this region passes 1% of the transmitted light, this is a Tval=100%/1%=100. And D=log (1/100)=2.0. This then is Dmax. The D range of the negative is Dmax-Dmin=2.0-0.0=2.0. So a scanner with a D value of } 2.0 would capture the tonal range of this neg. The tonal range of this neg is 100 tonal variations. Prints generally have D values between 1.7 and 2.0. So the same scanner would handle the neg and a typical print.
Let's say that the darkest area of the neg transmits 0.1% of the transmitted source light. This then yields Tval=100/0.1 =1000. And D=log 1000 = 3.0. Thus, this neg has 1,000 tonal variations in it from clear to black. If the neg has double the number of tonal variations (2,000) that would mean that the opacity is 2,000 and D=log 2000 = 3.3. At 4000 tonal range, D=3.6. Thus, each 0.3D equates to doubling the opacity range or halving or doubling the transmission value. Thus, if a scanner has a D rating of 3.0 (1000 tonal ranges) and the neg passes a corresponding 0.1% of the transmitted light, and another scanner has a D rating of 3.3 (2000 tonal ranges) and the neg passes 0.05% of the transmitted light--can you really tell one from the other?
The eye is more responsive to subtle changes in tonal variations in the white region of an image than in the dark ones. Thus, it is important to retain as much variability and rendition in the darkest areas of the negs (the highlights) but to do so without sacrificing detail in the shadows (clearer areas of the neg). This is done using the zone system and expansion and contraction of tonal range.
The response of the neg emulsion's exposure to light is not linear over the range of clear to full black. Its response curve is somewhat like a flattened S. The low exposure region is called the toe, which consists of the film base + fog density (Dmin). Moving up the curve is the straight line section (not exactly a straight line), and then to the shoulder. At this point, increasing amounts of light do not have corresponding quantitative amounts of change in density. Further exposure reaches saturation, or Dmax. At this point, no increase in exposure will change the density of the neg.
The ultimate goal is to maximize the straight line section and move Dmax as high as possible, without detrimentally affecting Dmin. I use contraction to accomplish this. The approach is to overexpose and underdevelop. With a neg, the rule of thumb is to expose for the shadows. If there is insufficient exposure in the shadow areas, there will not be any detail rendered.
Using Ilford FP4+ film, I rate it at ISO 80 (mfg=125). Development is done using stock developer diluted 1:1. It is used one time and discarded. Never replenish or try to refresh the developer. Development time will vary, depending on the tonal range of the scene. Here are some extreme examples of how this system works: (if you are offended by nude images, use the still life links. These following links are to fine art nudes and still life which were done using the zone system previously described)
[nudes] A. This shot illustrates a typical impossible shot. Inside the room, the exposure was about EV 3, the outside was dense fog with a starch white picket fence at EV 11. A scene like this with eight stops of variation would typically be shot to either render detail in the highlights (fog and fence) or in the inside room's details (shadows). To render shadow detail and retain highlight detail, the zone system was used as described. The image via this link is as-scanned on a UMAX Powerlook III of a 6x6cm negative using the transparency adapter. You can see the detail in the paint on the wall and can see the fence in the fog. http://www.photoweb.net/pw_gal_nude/pw_gal_nude_1/g_2.html
B. This shot shows great shadow detail despite the outside light creating a hot spot on the model's head. And the window on the left was rendered, despite the high level of light. http://www.photoweb.net/pw_gal_nude/pw_gal_nude_1/g_6.html
C. This shot shows a rather low contrast scene. There are no remarkable highlights. There is much material in shadow. Overexposure and ensuring at least two zones of exposure for the shadows and then overdeveloping N+1 achieved a perfectly printable neg. Again, this neg is shown as-scanned. http://www.photoweb.net/pw_gal_nude/pw_gal_nude_6/g_3.html
[Still life] A. high side lighting. http://www.photoweb.net/pw_gal_still/pw_gal_still_4/g_5.html
B. Low shadow lighting, highlights present. http://www.photoweb.net/pw_gal_still/pw_gal_still_4/g_6.html
C. Even lighting. There is actually more detail revealed than is showed in this web pix. http://www.photoweb.net/pw_gal_still/pw_gal_still_3/g_6.html
Non-web images of these negs are of course much better than those on-line. But I hope these illustrate the points of the zone system. I do think it can apply to TEM negs. The other point is to keep in mind the relationship of D values of scanners to what you are actually going to be scanning. If a scanner has sufficient resolution, and say a D rating of 3.2. Are you really going to be able to tell any difference using a scanner with a 3.4 or 3.5D at much higher cost? If you have a densitometer, check the D range of some of your negs. They are probably all less than 3.0. Maybe I'm wrong in this respect since I have little experience with TEM media. But the idea is go get the equipment you need for the job you need (the output destination and the use of the image) based on the actual media being scanned. Otherwise, there is a great opportunity to buy capability which will never be utilized.
Despite all the discussion of processing negs, as more digital capture and image processing products come out, things will change. There are rather simple ways to expand the contrast of an otherwise low contrast, poor neg. Likewise, there are ways to extract subtle detail from negs which have blown out highlights. More about this later.
gary g.
Reference: Adams, A. (1981). The negative. Boston: Little, Brown and Company. ISBN 0-8212-1131-5 (twelfth printing, 1992).
At 01:07 AM 4/12/2001, you wrote:
} Usualy to reduce contrast you under expose the film. Then develop it to } the disired density using a developer that generates low contrast. One way } to reduce contrast is to dilute you developer by a factor of 2, 4 or more } with water. It extends the developing time a good deal but it reduces the } contrast. You might also look at low contrast developers that work with } the film you are using. } } A few question on rec.photo.darkroom will get you more information than } you can handle and some of it will actually work. } } Gordon } Gordon Couger gcouger-at-couger.com } Stillwater, OK www.couger.com/gcouger
Good job, Gary! Gary's description of dynamic range is the clearest and most easily understood I have seen (and I have been looking for a good one!). I think I already understood most of what he said before he said it but I was having a devil of a time trying to articulate the concept to one of my staff. My only followup question is in regards to his statement near the end where he suggests some test the density of a TEM negative with a densitometer to see if they really go much above 3.0. I would like to encourage any one who has or will be measuring this for a typical and even finicky biological thin section TEM image to please post the info on the Microscopy listserver. Thanks.
} ------------------------------------------------------------------------ } 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 hesitated to make any response regarding this subject. } But--here goes anyway. Someone may find it useful. } } I think that the issue is not reduction of contrast but rather extension } of tonal range of the negative. That is what the zone system is } all about. I think that it is a fact that if the silver is not exposed, } there is not going to be any information gleaned from that } unexposed or too underexposed area. The approach I use for fine } art images is to overexpose and under-develop. It takes a LOT } of work to develop a complete system of ISO rating, development } and printing to achieve marketable prints. But it certainly can } be done. The goal is to minimize the time spent in the darkroom } doing printing. Ideally, the neg should print without any effort } on grade 3 paper. I only use Ilford archival matte fiber paper, so } extensions to RC papers may or may not directly apply. I also } do not use variable contrast paper. But the principles are extensible } I would think. } } Being all-digital with my SEM, I use realtime histogram feedback } to adjust gain and dynamic range. For TEM, I have no direct } experience. But I might suggest that the same photo techniques } for fine art neg work may apply to TEM negs. If you think that this } might be true, read on. Otherwise, nevermind. } } } A neg with bad dynamic range (huge extremes of EV or low contrast) is not } going to produce a very good scan. This is the realm of drum } scanners with 4.0-6.0D. Pixel resolution is subordinate to } D range in this case. If one is going to print a neg on paper, } that is one aspect of the problem. If the neg is to be output } to a magazine or printed page, that is another aspect. So the } crux of the matter is what the actual intended end use of the } neg is to be? If it is a nice print, OK. If it is a magazine or litho } output, these are 133lpi. Doing scans at 4000dpi only to } print them in a magazine at 133lpi is rather silly. One would } probably be better off just printing the neg and scanning at } 300dpi. } } But the 300 dpi is the effective dpi for the size of } the final image....not the original neg. So, if the neg is say } 3" x 4" and is to be printed at that same size, a 300 dpi scanner } should do the job (ignoring tonal range for the time being). } If the neg is to be printed at twice its physical size, then the } scanner has to have twice the resolution (600 dpi). And it } goes on from there. Thus, for most image output methods } on a printed page, 600-1200 optical dpi should be plenty of } resolution. If you agree with this discussion at this point, } then the resolution factor should no longer be an issue. } What remains is D range. } } (Note that commercial images are created at high resolution } to accommodate manipulation and merging with other images. } This way, image quality can gradually be reduced without } affecting the final result.) } } But the negative is the key item in all of this discussion. A } bad neg is not likely to do anyone much good. So the challenge } is to get a good neg at the get go. Here we go, back to the } zone system. The idea is to use the neg as a non-linear } image capture media for data which may span a wide } dynamic range. And in so doing, be able to output (print } or scan) this information with fidelity and minimal effort. } This drives D value. } } An unexposed, developed neg will have a transmission of 100%. } It is actually slightly lower due to absorption by the medium, } emulsion and residual anti-dispersion coating. But at 100% } transmission, the neg would have an opacity of 100%input/100%output } or 1.0. Since D=log1/T, the D value for the unexposed neg } is log 1/1=0.0. This then is Dmin. The area with the most } exposure (darkest region) will pass the least amount of } transmitted light. If, for example, this region passes 1% of } the transmitted light, this is a Tval=100%/1%=100. } And D=log (1/100)=2.0. This then is Dmax. The D range } of the negative is Dmax-Dmin=2.0-0.0=2.0. So a scanner } with a D value of } 2.0 would capture the tonal range of this } neg. The tonal range of this neg is 100 tonal variations. } Prints generally have D values between 1.7 and 2.0. } So the same scanner would handle the neg and a typical } print. } } Let's say that the darkest area of the neg transmits 0.1% } of the transmitted source light. This then yields Tval=100/0.1 } =1000. And D=log 1000 = 3.0. Thus, this neg has 1,000 } tonal variations in it from clear to black. If the neg has double } the number of tonal variations (2,000) that would mean } that the opacity is 2,000 and D=log 2000 = 3.3. At 4000 } tonal range, D=3.6. Thus, each 0.3D equates to doubling } the opacity range or halving or doubling the transmission } value. Thus, if a scanner has a D rating of 3.0 (1000 tonal } ranges) and the neg passes a corresponding 0.1% of the } transmitted light, and another scanner has a D rating } of 3.3 (2000 tonal ranges) and the neg passes 0.05% of } the transmitted light--can you really tell one from the other? } } The eye is more responsive to subtle changes in tonal } variations in the white region of an image than in the } dark ones. Thus, it is important to retain as much variability } and rendition in the darkest areas of the negs (the highlights) } but to do so without sacrificing detail in the shadows (clearer } areas of the neg). This is done using the zone system } and expansion and contraction of tonal range. } } The response of the neg emulsion's exposure to light is not linear } over the range of clear to full black. Its response curve is somewhat } like a flattened S. The low exposure region is called the toe, which } consists of the film base + fog density (Dmin). Moving up the curve } is the straight line section (not exactly a straight line), and then to } the shoulder. At this point, increasing amounts of light do not } have corresponding quantitative amounts of change in density. } Further exposure reaches saturation, or Dmax. At this point, } no increase in exposure will change the density of the neg. } } The ultimate goal is to maximize the straight line section and } move Dmax as high as possible, without detrimentally affecting } Dmin. I use contraction to accomplish this. The approach is } to overexpose and underdevelop. With a neg, the rule of thumb } is to expose for the shadows. If there is insufficient exposure } in the shadow areas, there will not be any detail rendered. } } Using Ilford FP4+ film, I rate it at ISO 80 (mfg=125). Development } is done using stock developer diluted 1:1. It is used one time } and discarded. Never replenish or try to refresh the developer. } Development time will vary, depending on the tonal range of the } scene. Here are some extreme examples of how this system works: } (if you are offended by nude images, use the still life links. These } following links are to fine art nudes and still life which were done using the } zone system previously described) } } [nudes] } A. This shot illustrates a typical impossible shot. Inside the room, } the exposure was about EV 3, the outside was dense fog with } a starch white picket fence at EV 11. A scene like this with } eight stops of variation would typically be shot to either render } detail in the highlights (fog and fence) or in the inside room's } details (shadows). To render shadow detail and retain highlight } detail, the zone system was used as described. The image } via this link is as-scanned on a UMAX Powerlook III of a } 6x6cm negative using the transparency adapter. You can } see the detail in the paint on the wall and can see the fence in } the fog. } http://www.photoweb.net/pw_gal_nude/pw_gal_nude_1/g_2.html } } B. This shot shows great shadow detail despite the outside light } creating a hot spot on the model's head. And the window } on the left was rendered, despite the high level of light. } http://www.photoweb.net/pw_gal_nude/pw_gal_nude_1/g_6.html } } C. This shot shows a rather low contrast scene. There are no } remarkable highlights. There is much material in shadow. } Overexposure and ensuring at least two zones of exposure } for the shadows and then overdeveloping N+1 achieved } a perfectly printable neg. Again, this neg is shown as-scanned. } http://www.photoweb.net/pw_gal_nude/pw_gal_nude_6/g_3.html } } } } [Still life] } A. high side lighting. } http://www.photoweb.net/pw_gal_still/pw_gal_still_4/g_5.html } } B. Low shadow lighting, highlights present. } http://www.photoweb.net/pw_gal_still/pw_gal_still_4/g_6.html } } C. Even lighting. There is actually more detail revealed } than is showed in this web pix. } http://www.photoweb.net/pw_gal_still/pw_gal_still_3/g_6.html } } } } Non-web images of these negs are of course much better than } those on-line. But I hope these illustrate the points of the zone } system. I do think it can apply to TEM negs. The other point } is to keep in mind the relationship of D values of scanners to } what you are actually going to be scanning. If a scanner has } sufficient resolution, and say a D rating of 3.2. Are you } really going to be able to tell any difference using a scanner } with a 3.4 or 3.5D at much higher cost? If you have a } densitometer, check the D range of some of your negs. They } are probably all less than 3.0. Maybe I'm wrong in this } respect since I have little experience with TEM media. } But the idea is go get the equipment you need for the job } you need (the output destination and the use of the image) } based on the actual media being scanned. Otherwise, there } is a great opportunity to buy capability which will never be } utilized. } } Despite all the discussion of processing negs, as more } digital capture and image processing products come out, } things will change. There are rather simple ways to expand } the contrast of an otherwise low contrast, poor neg. Likewise, } there are ways to extract subtle detail from negs which have } blown out highlights. More about this later. } } gary g. } } } Reference: } Adams, A. (1981). The negative. Boston: Little, Brown and Company. } ISBN 0-8212-1131-5 (twelfth printing, 1992). } } } } At 01:07 AM 4/12/2001, you wrote: } } } Usualy to reduce contrast you under expose the film. Then develop it to } } the disired density using a developer that generates low contrast. One way } } to reduce contrast is to dilute you developer by a factor of 2, 4 or more } } with water. It extends the developing time a good deal but it reduces the } } contrast. You might also look at low contrast developers that work with } } the film you are using. } } } } A few question on rec.photo.darkroom will get you more information than } } you can handle and some of it will actually work. } } } } Gordon } } Gordon Couger gcouger-at-couger.com } } Stillwater, OK www.couger.com/gcouger
-- 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)
I tried sending this earlier, but seems to have not made it.
The 840 has some pots (potentiometers) on one of the circuit boards that are used to adjust out any image shift during stigmation adjustments. They probably adjust some bias voltages, or some offsets, in the stigmator circuits. They can be adjusted so there is no image shift.
If the pots can't eliminate the image shift, then a component has probably gone bad, rather than just drifted a bit. If you have the manual with the schematics, they should be identified in there. If you don't have the manual, I might be able to find them in ours. Let me know.
The 18th Annual NESM Woods Hole Symposium will be held May 11-12th, at the Marine Biological Lab at Woods Hole, MA.
This meeting is supported by members of the Connecticut Microscopy Society (CMS), the Metropolitan Microscopy Society (MMS), and the New York Society of Experimental Microscopists (NYSEM).
Pre-registration is encouraged and is a must if you plan to attend the Friday night dinner. Inquiries re: registration for this meeting should be directed to Mary McCann (617) 484-7865 or by email: mccanns-at-tiacc.net. Advance regis- tration including dinner on May 11th is $40.00 for NESM members, and $55.00 for non-members (this cost includes a one- year membership to NESM).
PLEASE NOTE: ADVANCE REGISTRATION MUST BE RECEIVED no later than MAY 4th!!! Registrations received after May 4th will NOT include dinner.
Friday, May 11th begins at Noon and consists of 2 sessions with an afternoon coffee break. Following the presentations, a cocktail hours and dinner will commence in the Swope Center.
Saturday, May 12th, NESM will present a symposium on Remote Access Microscopy. Afterwards, there will be commercial exhibits and posters on display in the Swope Center. Presentation of Poster and Photos-As-Art Awards and Door Prizes will follow. This year lunch at the Swope Center will be optional; those interested will pay $16.00. After lunch, two 45-minute tours of the Marine Resource Center will take place.
NESM welcomes new members to the Society! Please join us for this most enjoyable meeting.
Peggy Sherwood, Corresponding Secretary NESM -- Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 50 Blossom Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) sherwood-at-helix.mgh.harvard.edu
You have a really good point for discussion about actual available exposure time. Real world nudes and still lifes can be for several seconds exposure. What is it for a typical TEM neg? I don't know. I suspect the same drift problems occur in TEM as they do in SEM. So I too like to get the shot as quickly as possible yet with minimum noise. So it is a tradeoff of pixel density and pixel dwell time. Some original loss can be made up later using the computer. With a digital active scan and image capture system synchronized to 60 Hertz, a SEM shot at 3000x3200 pixels takes 96 seconds at 10uS dwell time. This works fine. But for a TEM which may not have sync to line frequency, I can see that image shift is a big problem. Are TEM cameras and scanning routinely synchronized to line frequency?
I used Diafine and Accufine back in photojournalism days to push process TriX to ISO1600 or beyond. The job was to shoot basketball and football games without strobe. The reason was to gain even lighting and high contrast shots. Since the pix were physically small, the higher grain was not a big issue.
Your Diafine approach to TEM negs sounds like a solid method. Indeed, the key is to get the shot and make a print with minimum amount of tweaking (burning, dodging, etc.). If one were to only have to make one print ever from one neg, it would not matter all that much. But if more than one print is or will be made, reproducibility is a tough issue with a less than perfect neg.
I see a similarity in your approach and what I was talking about. Your procedure limits the development time of the neg. But while it limits the time for dense areas, it limits the time for the whole neg too. The procedure I described ensures that as much light information is captured on the film, but extends the tonal range of the film by reducing the strength of the developer and the development time.
I already have de-rated the speed of the film in this procedure. It may be that doing this with TEM negs makes the resulting absolute speed too slow. What is the rated ISO speed of some TEM media in use today?
gary g.
At 01:56 PM 4/13/2001, you wrote: } Reply to: RE: Film Processing & dynamic range & scanners (longish) } I saved your extensive info about negs. I would not use Diafine for } regular scenic photos--too low in contrast. However, the manner in which } split developers achieve this is to LIMIT the amount of developer } available to "dense" negative areas. I only use it for contrasty TEM } negs. that I routinely print on #2 or #3 paper , whichever is more } pleasing. By having easily retrievable info in all of a negative, less } time should be needed by any final printing method. The developer has a } long tank life and works at room temperature. An extreme test of TEM negs } made at 4 f/stops equivalent underexposure still printed nicely on #3 } grade paper. } Of course I used a very long 12 mimutes in each half of the split } developer, but this can allow short exposure on specimens with a } "drifting" problem and save an expensive experiment. } As primarily an old "wet" printer, I enjoyed your thorough } explaination and have heard of the zone system and the great A. Adams. I } like to get good results with little effort, thats all. If you ever want } a copy of the Diafine Co.'s explaination of it's product, I can send you } one. Nearly all of my published photos in Ultramicroscopy were done with } this stuff, including a "lucky" cover photo on Ultra. 25 (1988) 351-354. } } Bernie Kestel E-mail: {kestel-at-anl.gov} } Materials Science Division } Argonne National Lab } 9700 So. Cass Ave., } Argonne Il., 60439 } Gary Gaugler wrote:
Has anyone had experience in determining the concentration of particles in a solution by counting on EM grids in a similar way to using a haemocytometer? Is it possible to count the particles in a defined number of grid squares then calculate back to the area of the grid and the amount of solution which was applied and allowed to dry down?
Dear Richard, I can forsee one difficulty. The particles may not be deposited uniformly due to a number of factors: 1) Evaporation of the applied drop of specimen will not be uniform, so the particles could be dragged in toward the grid center as the edges evaporate (or they could be preferrentially deposited toward the edges if they are hydrophobic). 2) The grid surface may not be flat, causing particles to deposit preferrentially in the centers of the grid squares--if these are lower--or near the grid bars. 3) The particles could be preferrentially deposited either on top of the grid bars or on the open areas, which could look like uniform deposition, but would give erronious quantitation. Of course, one could test this by depositing a known amount of suspension of a known concentration of the kind of particles to be measured, then seeing if the grid was covered uniformly and if the calculation gave the correct result. Good luck. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
I asked if xylene was the best solvent for cleaning immersion oil off a non-immersion objective. I recalled in my brief training that xylene was safe but alcohols dissolve the more common lens adhesives (I could be wrong and have this reversed). Here is the array of responses I received. The diversity may interest you. THANKS to all for your advice.
1 We regularly use xylene with a cotton tipped applicator to clean our lenses and remove immersion oil. This was recommended to us by the supplier of our lenses and microscopes for a number of reasons. The primary reason is that xylene effectively 'cuts' the oil, without damaging the lens. Acetone will affect the lens coating, and Isopropyl alcohol only rinses the oil without effectively removing it. Hope this helps
2 You are lucky that the lens elements did not dislodge. Use alcohol on lens paper. Never use xylene or toluene; these solvents can dissolve the cement used to seat the various lens elements.
3 I would recommend Sparkle Glass Cleaner. I only use solvents as a very last resort (unless you really, really do want to upgrade) The cements are soluble in most solvents. With intractable grime, I use a cotton tipped applicator moistened with either xylenes or toluene and shake all excess off. The tip must be moist, not wet or dry and make single passes until the grime come off. Only enough solvent on the applicator to dampen the surface, not enough to wet.
4 For day to day cleaning of lenses, including oil on the 40X. (I can't believe this is the first time! The graduate students and Post Doc's drag the40X through the oil at least twice a week. I have given them repeated instructions but they seem intractable) Invert an ocular and examine the surface for cleanliness. If it is clean, don't clean it. Dust off any loose debris. Check the lens again. Moisten a cotton tipped applicator in Sparkle and wipe the lens 1 swipe with a rolling action to present a new surface and lift off any grit. Discard. Take a dry applicator and remove the film. Check the lens with an inverted ocular to see if it is clean. Do no more than is necessary. There has been an ongoing rampage on the list about lens cleaning.... to solvent, not to solvent... lens tissue, not to lens tissue etc.. I can append you a couple, and a microscope maintenance handout if you would like. It will be a bit long about 15 pages (38K).
5 The care instructions for my immersion oil suggest cleaning with a soft cloth or lens tissue (no Kimwipes) moistened with ether/alcohol (7:3) or xylenes. My microscope manuals recommend removing finger prints using alcohol. Sounds like you're good to go.
6 We use 70 % isopropanol to clean emersion oil off of our lenses.
7 Cleaning with xylene is a little drastic. If the lens eventually gets xylene in behind the cement it could do some damage and you would have to send it in to Nikon for repair. I use Kodak lens cleaner and some lens cleaning tissues (not regular Kim wipes) to get the oil off, constantly checking them under a dissecting scope to make sure that it is all removed. It works well on some expensive lens that we have here.
8 We use Green Soap from the pharmacy. It works the best with ultra pure water.
9 Actually, you've got your solvents reversed; alcohol is usually safer. But safer still is diluted detergent "Joy" or similar, followed by water. Use alcohol only if that doesn't work. And fumes from xylene, toluene, etc. are best avoided.
10 Back in the pre PC (chemical) days, toluene was a recommended } solvent for cleaning lenses of immersion oil. Now, we at National Steel, } wipe off excess oil with lens tissue and then clean the lens with Kodak lens } cleaner solution.
Ritchie, What you are describing is a stigmator drive that is out of calibration. If you look on the schematic for the stigmator drive, you should see the X(or Y) control that goes to an op-amp that in turn feeds a voltage divider which in turn controls a different coil in the stigmator coil assembly. Each part of this voltage divider has its own trim pot for calibration purposes. Simply rock the X stigmator control back and forth while adjusting each pot for minimum image shift. Then, increase your mag & repeat. You should be able to get all of the shift completely out. Good luck!
Gary M. Easton, Pres. Scanners Corporation SEM/EDS/IMAGING Sales & Service
----- Original Message ----- } From: "Ritchie Sims" {r.sims-at-auckland.ac.nz} To: {microscopy-at-sparc5.microscopy.com} Sent: Friday, April 13, 2001 2:12 PM
Thanks very much to all those who responded to my post.
My hands didn't bleed at all.
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
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At 12:53 PM -0400 4/10/01, Ronald Anderson wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I am sending this message again because it was blocked by a list server filter - presumably because 'home' is part of your e-mail address.
Malcolm
-------- Original Message --------
Please reply directly to:
} From: {hmskaug-at-tartarus.uwa.edu.au} } } Subject: TEM } } } Hi, } } I am doing some preperation work for an assignment, and have a question. } } If a fresh unfixed piece of brain tissue (weighing approx. 1 gram) is } received in a diagnostic electron microscopy laboratory, how would I } proceed with the specimen? The brain is taken in surgery within the last 5 } min. } } If I was asked to carry out a rapid viral diagnosis on the tissue, what } precautions should I take, and how would I proceed? } I read that prions are able to survive fixation. What then? } } A respond is very much appreciated. } } Thank You ! } } Sincerely, } } Hege Skaug
Greg Erdos Assistant Director Biotechnology Program Ph. 352-392-1295 University of Florida Fax 352-846-0251 PO Box 118525 Gainesville, FL 32611 http://www.biotech.ufl.edu/~emcl
To all colleagues who supply standards for EDS work I need a standard sample of known thickness to check the accuracy of my TEM/EDS system during standardless quant. I need to include light elements (Z {11) as I have a Moxtek SUTW window on my detector. I guess an amorphous glass sample that contains both nitrogen and oxygen with with a very thin layer of carbon on it to prevent charging would do the trick. Can anybody out there supply one? Thanks.
Alan Fox
Professor Alan G. Fox BSc PhD CEng FIM Director, Center for Materials Science and Engineering Naval Postgraduate School Monterey California 93943 USA
This is an apology written in embarassment for some of my private comments about EDX systems that recently got broadcast to the list. I had too-quickly responded offline to a posting and was chagrined to find my comments passed along without the balanced context I would have put them in if I'd been more sensitive to the probability of dissemination. It's too late to rephrase them but I want to emphasize that my recent demos with Oxford, Noran, PGT and EDAX were all highly positive experiences and I came away believing that all are solid and supportive companies offering world class instrumentation. Any system will have many pros and a few cons; our final choice will rest simply on which has a bigger cluster of the former than the latter for our particular set of applications. The cons I had mentioned were subjectively-perceived blips within the significent strengths that each system offers. While cons can serve to pare down the choice, once it's made, as I've discovered through discussion with reference users representing all four companies, users across the board (at least all those I've spoken to) are happy with whichever system they purchased.
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)
There is a decent test sample out there for checking and monitoring your EDS system called the NiOx sample. One place that you can find it is at Ted Pella. Here is the web site for that product. It has further information on it. http://www.tedpella.com/calibrat_html/TEM7.htm
I am not sure what you mean by the accuracy of your EDS system. You will need to calibrate it and determine whether you are having problems with icing. You can do that with the NiOx sample. The literature that comes with the sample or that you can get from Ted Pella will tell you how to do this. It is also tells you how to monitor your system's performance over time.
If you want to check for N2, just get some Hexagonal BN, crush it between two glass slides, and collect it on a carbon coated grid. You will easily find thin particles.
You hit on a couple of problems with your request. You will have a difficult time ascertaining the thickness of any sample accurately, but with glass, you will only be limited to doing it with EELS or contamination spots. With glass, the composition can change under the beam as elements particularly alkali elements diffuse out of the irradiated area. The composition of glasses can change depending on the depth from the surface and so where you are can make a big difference. I do not know where you would get a glass with a N2 concentration. You can also cause the glass to soften in the beam in very thin areas if care is not taken. Higher accelerating voltages help here tremendously.
I have found that frequently I do not need to coat my glass samples in a 200 keV TEM. I did have to do it when I used a 100 keV machine. For cross section samples, I started using Si blanks as half of the sample and it seems to have eliminated heating and charging problems.
-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: Alan Fox [mailto:fox-at-nps.navy.mil] } Sent: Monday, April 16, 2001 12:50 PM } To: MS listserver } Subject: Standard sample for EDS quant } } } -------------------------------------------------------------- } ---------- } 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 } } -------------------------------------------------------------- } ---------. } } } To all colleagues who supply standards for EDS work } I need a standard sample of known thickness to check the } accuracy of my TEM/EDS system during standardless quant. I need to } include light elements (Z {11) as I have a Moxtek SUTW window on my } detector. I guess an amorphous glass sample that contains } both nitrogen } and oxygen with with a very thin layer of carbon on it to prevent } charging would do the trick. Can anybody out there supply one? Thanks. } } Alan Fox } } } Professor Alan G. Fox BSc PhD CEng FIM } Director, Center for Materials Science and Engineering } Naval Postgraduate School } Monterey } California 93943 } USA } } Tel (831) 656 2142 (work) } (831) 657 9239 (home) } Fax (831) 656 2238 } } }
May I suggest that we consider some ground rules regarding postings of summaries of replies to questions? It happens on occasion that replies which are assumed to be made in private get posted publicly, sometimes to the embarrassment of those doing the replying. This is, I'm sure, never done with any bad intent and is usually harmless , but can nonetheless be a little disconcerting.
I have posted summaries myself without thinking of the possible consequences, so I'm not pointing any fingers. It's something that's easy to forget about until you get caught yourself.
I would suggest as starting points that: 1) questioners always make clear their intent to post a summary, 2) that the identities of the repliers be removed from summaries (this is often done anyway), and 3) that all who reply to a question indicate if they want their replies to be kept private.
Anyone else have any thoughts on this matter?
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/
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I would need to prepare DNA for TEM using spreading/shadowing in Cytochrome C films. My pilot experiments using just plasmid DNA and the Lang & Mitani method (Biopolymers, 9, p.373, 1970) worked rather poorly and I would like to hear from people with some experience in this method. What are the critical points here? Purity of water/chemicals, Cyt C concentration, time? Any hints would be appreciated.
Do I understand you believe this is CJD or prion infected tissue? If so several methods of disinfection have been suggested (using the word disinfect loosely). No one is sure exactly what will inactivate this prion and the procedure has been disgusted on Histonet for routine pathology giving the latest suggested methods. I can attempt to forward some of that information to you along with the website address. Let me know if you would like the information. Most of us are alittle frightened of this one as it can take years to manifest. Pamela A. Marcum Histology/Microscopy Product Development Manager 400 Valley Road Warrington, PA 18976 Phone: 800-523-2575 Ext 167 215-343-6484 Ext 167 Fax: 215-343-0214 E-mail: pmarcum-at-polysciences.com
-----Original Message----- } From: Greg Erdos [mailto:gwe-at-biotech.ufl.edu] Sent: Monday, April 16, 2001 9:17 AM To: Microscopy-at-sparc5.microscopy.com
Please reply directly to:
} From: {hmskaug-at-tartarus.uwa.edu.au} } } Subject: TEM } } } Hi, } } I am doing some preperation work for an assignment, and have a question. } } If a fresh unfixed piece of brain tissue (weighing approx. 1 gram) is } received in a diagnostic electron microscopy laboratory, how would I } proceed with the specimen? The brain is taken in surgery within the last 5 } min. } } If I was asked to carry out a rapid viral diagnosis on the tissue, what } precautions should I take, and how would I proceed? } I read that prions are able to survive fixation. What then? } } A respond is very much appreciated. } } Thank You ! } } Sincerely, } } Hege Skaug
Greg Erdos Assistant Director Biotechnology Program Ph. 352-392-1295 University of Florida Fax 352-846-0251 PO Box 118525 Gainesville, FL 32611 http://www.biotech.ufl.edu/~emcl
To all colleagues who supply standards for EDS work I need a standard sample of known thickness to check the accuracy of my TEM/EDS system during standardless quant. I need to include light elements (Z {11) as I have a Moxtek SUTW window on my detector. I guess an amorphous glass sample that contains both nitrogen and oxygen with with a very thin layer of carbon on it to prevent charging would do the trick. Can anybody out there supply one? Thanks.
Alan Fox
Dear Alan, I can't get you a standard with all the qualifications you want, but I do have some standards prepared by Chuck Fiori. The matrix is a lithium borate glass, and there are several blocks with various elements evenly dispersed in the matrix. The down side is that you will have to melt the glass, prepare thin specimens--by blowing with a platinum straw--break off pieces from the bubble, and put them on (or in) a suitable grid. I have found that folding grids work well. Of course, you will have to measure the thickness after you prepare the specimen. Anyway, it is amorphous glass with light elements, including oxygen. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
We are trying out the suggestion of using Nile Red to increase fluorescence. This seems very promising at this stage.
Ian
Ian Hallett HortResearch Mt Albert Research Centre Private Bag 92 169 Auckland, New Zealand Fax 64-9-815 4201 Telephone 64-9-815 4200 EMail ihallett-at-hortresearch.co.nz
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We are looking at purchasing a stereo microscope with fluorescence capabilities primarily to look at GFP fluorescence in plant specimens. Does anyone have any particular comments on the relative merits of the systems produced by the major microscope manufactureres (Leica, Nikon, Olympus and Zeiss). We also have a dichotomy amongst users on whether to provide film or digital cameras for recording - my own preference is more on the digital side but again has anyone any comments or suggestions.
Thanks
Ian
Ian Hallett HortResearch Mt Albert Research Centre Private Bag 92 169 Auckland, New Zealand Fax 64-9-815 4201 Telephone 64-9-815 4200 EMail ihallett-at-hortresearch.co.nz
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This is a cry for help on behalf of an unwell EDS detector that has been having some problems. It has been behaving very strangely lately but does not know who to turn to....
For the past six months it has been suffering short-lived episodes of low energy tails accompanied by some visible peak broadening most apparent at the low energy end of the spectrum. I am aware of previous discussions about possible causes of tails and peak broadening but I think the unusual thing in this case is that the problem only lasts for the first few minutes after the SEM beam is turned on and the detector is first exposed to X-rays for the day. It then gradually goes away and all aspects of the detector's behavior return to normal again. The tails won't return again that day once the SEM has been used.
This is a 10mm2 127eV Noran "Pioneer" detector with an atmospheric thin window hooked up to a Voyager IV with digital pulse processing. We leave the whole system running all of the time. It is attached to a Jeol SEM and we normally use its airlock to change samples which means that the crystal/window do not see daylight/atmospheric pressure very often. We routinely use the "slowest" highest resolution pulse processor settings. Except for this short-lived problem everything else about the system is as good as it ever was when we first installed it and ran the pulse processor setup routines a few years ago. As far as we can tell there is no *detectable* buildup of ice inside the detector (no phantom oxygen peaks etc) and there is only a small amount of visible oil contamination on the window.
It takes 12 hours or more of idleness before the tails re-appear, where then it can take 10 to 20 minutes exposure to X-rays for them to go away again. It takes less time for the tails to go away if the beam current is temporarily ramped up high enough to "saturate" the system with enough counts to approach 100% dead time. The only other unusual features that we have observed after the detector has been idle for a while are that the idle dead time indication fluctuates wildly from zero to ~50% or more from reading to reading (two second sampling periods) instead of the usual 10-20 percent, and the idle Detects and Converts are only a few counts/sec instead of the normal several tens of counts/sec. After a good dose of X-rays these parameters return to their normal idle values again also.
There is another piece to this puzzle. Last week we gave the detector a "photon enema" with a flashlight while examining the window and surrounds for anything unusual and that had the same effect in curing the problem as using a high beam current to generate X-rays.
So does anybody out there have some clues about this -- a cause, a cure, or even the physics of it? I've got the impression that it must be something electronically weird in relation to the crystal, such as some sort of an excess charge accumulation when it is idle for a long time which then gradually dissipates when the crystal starts responding to X-ray photons again? Could this effect occur if the bias was just slightly wrong by a few volts? How about "leakage current" or FET problems? I haven't got the foggiest idea really because the zero position and peak energies are always ok.
Very strange?
Concerned, Australia.
Arthur Day, Electron Microscope Unit Phone: 61-2-9717-3457 Ansto Materials Division Fax: 61-2-9543-7179 PMB 1, Menai (Sydney), NSW, 2234 Email: ard-at-ansto.gov.au Australia www: http://www.ansto.gov.au/
Ian, I have no specific comments with the new breed of fluorescent stereo's. However, in going through this a decade ago with "conventional" stereos, I found that different scopes though all well designed ranked differently with different specimens. The objects that we biologists stare at are odd optically. I suggest that you get demos and inspect the favorite objects in your departement that will be viewed under the scope. I found one stereo whose darkfield far outshone the others for arabidopsis roots, though there is no "rational" reason for this, and nothing to say that this scope is "better" in general.
A key feature to insist on is a shunt with 100% of the light to the phototube. In the old days most stereos did not do this, perhaps with fluoresecence things have changed, but you want all the light going for the captured image, whether it is film or pixels.
For image capture, going directly to a color slide can be handy. On my stereo, we trade off between video and film quite a lot. Our 35 mm camera back is no big deal (that is, just a cameraback on a tube) and adding that capability did not add much to the cost. You can of course buy much fancier and expensive 35 mm controllers but my point is you don't have to.
Hope this helps, Tobias
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Question: How long should it take for agar to conjeal after I pour it into a petri dish?
Dear Billone, Way back when I was pouring agar for immunodiffusion plates, I found that it congealed almost immediately. I had some trouble keeping it liquid to get a good smooth surface. Since that applies to the specific concentration of agar I used (I forget, but I think it was ~1%) and the temperature it was heated to (~40 C, I think), and since that was before global warming, YMMV. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
Dear Ian, The recommendation from Tobias to compare before buying is always sound. We've been using a Leica MZ-12 epi-fluorescent stereomicroscope for about 2 years. GFP is only part of its use, we do a lot of microinjection and of late have been using it with vitally stained zebrafish larvae. There are a number of practical considerations. All of the mfrs. originally had limited ability to change wavelengths, the original Leica required removing the eyepieces and camera adapter as a unit, then removing screws to withdraw the filterset. Their current FL-III system has a rotating ring holding several filtersets. Filter combinations may be removed and inserted easily with it. We shuffle about 7 different sets between the filter ring, which holds about 6 (and we keep one open for brightfield). Don't ever think you will never need more than just GFP.
Adding a camera tube and fluorescent lamp housing makes the scope head very heavy. We have ours on a double arm boom (Diagnostics Inst.) to maneuver micromanipulators beneath it, and it still vibrates when you touch it. The focus must be kept tight. I just bought a lab jack stand to support our samples and to provide fine focus. Pay attention to flex in the manner in which the scope head is attached to the focus mechanism, and to the boom.
With fluorescence, consider the NA of the lenses, as that will determine whether the fluorescence is visible. We use a .8X/.05 NA for general use and long working distance with microinjection equipment. But the 1X/.125 allows us to see GFP transfected neuronal processes. A new 1.6X/.2 lens provides several times greater intensity allowing some projects to be done quickly with the stereoscope instead of having to be transferred to a compound scope.
I'd be interested in your digital camera decision. The weight issue limits what we can install. Weve been sticking with a no-name single chip color camera, which is very lightweight. The resolution is fair, and it is surprising light sensitive. The users who prefer it are viewing the color output on a passthrough monitor, then capture monochrome. The color is essential when doing multi-label injections. Color capture resolution from it is dreadful, but the monochrome is decent resolution. Other color cameras have been too heavy, inducing big vibrations when you touch the focus, or have not been sensitive enough. A Nikon Coolpix 990 is terrific for brightfield, but lacks sensitivity for fluorescence. You can get pretty good at guesstimating exposure times for 1-8 second exposures of dim fluorescence. But, the chip noise isn't worth it. And, if you need real-time imaging of low-light images for any reason (there are several reasons you might) then the Coolpix won't work. The final straw with the Coolpix, and some other cameras, is that we use some far-red dyes, like DiD, to which the Nikon is blind at working concentrations.
Regards, Glen
} } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } We are looking at purchasing a stereo microscope with } } fluorescence capabilities primarily to look at GFP fluorescence in } } plant specimens. Does anyone have any particular comments on } } the relative merits of the systems produced by the major } } microscope manufactureres (Leica, Nikon, Olympus and Zeiss). } } We also have a dichotomy amongst users on whether to provide } } film or digital cameras for recording - my own preference is more on } } the digital side but again has anyone any comments or } } suggestions. } } } } Thanks } } } } Ian } } } } } } Ian Hallett } } HortResearch } } Mt Albert Research Centre } } Private Bag 92 169 } } Auckland, New Zealand } } Fax 64-9-815 4201 } } Telephone 64-9-815 4200 } } EMail ihallett-at-hortresearch.co.nz } } }
-- Glen MacDonald UW Core for Communications Research Virginia Merrill Bloedel Hearing Research Center Box 357923 University of Washington Seattle, WA 98195-7923 glenmac-at-u.washington.edu (206) 616-4156
Alan - The National Institute of Standands and Technology (NIST) sells a well-characterized Mg-Si-Ca-Fe-O glass thin film which is intended as a standard for EDS calibration in the TEM. The glass is supported by a 20 nm carbon thin film and its thickness has been measured by profilometry. I routinely use this standard along with other mineral standards for EDS calibration in the TEM and over the years have found it to be a very good standard indeed.
NIST can be contacted at (301)975-6776.
Dave
Dave Joswiak Dept. of Astronomy, 351580 University of Washington Seattle, WA 98195 (206)543-7702
On Mon, 16 Apr 2001, Alan Fox 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 } -----------------------------------------------------------------------. } } } To all colleagues who supply standards for EDS work } I need a standard sample of known thickness to check the } accuracy of my TEM/EDS system during standardless quant. I need to } include light elements (Z {11) as I have a Moxtek SUTW window on my } detector. I guess an amorphous glass sample that contains both nitrogen } and oxygen with with a very thin layer of carbon on it to prevent } charging would do the trick. Can anybody out there supply one? Thanks. } } Alan Fox } } } Professor Alan G. Fox BSc PhD CEng FIM } Director, Center for Materials Science and Engineering } Naval Postgraduate School } Monterey } California 93943 } USA } } Tel (831) 656 2142 (work) } (831) 657 9239 (home) } Fax (831) 656 2238 } } } }
Hi, Your agar should be setting up almost immediately. I usually use a concentration of 1-2% for my purposes. One imperative point is that the agar needs to be completely dissolved. This may sound like a trivial comment, but I know that when I first started working with agar I made the mistake of not heating the solution hot enough for a long enough time and wondered what was wrong. If you have access to an autoclave, autoclave it for about 15 minutes. If not, heat it on a hot plate (preferably with a stir bar, but you can swirl it periodically if you don't have a stir bar) until the agar is completely dissolved and you have a nice, clear solution. Be careful, because agar can suddenly start to boil and be up and over the top of your flask before you realize it. Let me know if you need any more help. Good luck, Kristen
At 04:38 PM 4/16/01 -0500, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Kristen A. Lennon, Ph.D. Department of Plant Pathology 351 Bessey Hall Iowa State University Ames, IA 50011 515-294-8854 kalen-at-iastate.edu
In a message dated 04/17/2001 8:23:01 AM US Mountain Standard Time, glenmac-at-u.washington.edu writes:
{ { Adding a camera tube and fluorescent lamp housing makes the scope head very heavy. We have ours on a double arm boom (Diagnostics Inst.) to maneuver micromanipulators beneath it, and it still vibrates when you touch it. The focus must be kept tight. I just bought a lab jack stand to support our samples and to provide fine focus. Pay attention to flex in the manner in which the scope head is attached to the focus mechanism, and to the boom. } }
Glen's message is full of great advice. Regarding the point mentioned above, this is indeed crucial as you start stacking photo tubes, ergo heads, cameras, etc. onto the stereomicroscope. You can easily overwhelm the load limits of the focus drive and the gearing, and the scope will start to "droop" so that you have to continually bring the scope back into focus.
I don't know about other scopes, but Leica has a tension upgrade kit that can be installed in the focus drive unit for these situations. It makes the focus much tighter. This upgrade is for the MZ FL III which is configured (usually) on a transmitted light base with a focus post and focus drive attached to the base. As far as I know, there is no such kit if you decide to put the scope on a boom stand or a swinging arm stand. In that case, you would have to deal with the tightness of the articulated joints on the stand.
Anyway, if you order the MZ FL III stereofluorescence scope in the usual configuration, Leica has a cure for the "droopy scope" syndrome if it gets too heavy. For details on the upgrade you can contact Leica Customer Service at 1-800-248-0123. Or contact me and I will put you in touch with our Service Engineers who have installed several of the tension upgrade kits. One qualifier: installing the kit and the springs requires disassembling the focus drive unit. A factory trained service guy should definitely do this.
Good luck!
Best regards,
Bob Chiovetti GTI Microsystems, Inc. Leica Exclusive Regional Dealer Southwestern United States
I'm going out on a limb here, because it is not a particular area of expertise of mine, but I've not seen it in the discussion. Experts in microscopy of radiation sensitive materials are much more familiar with these issues (Linn Hobbs - can you help us here?)
I'm surprised that no-one has pointed out a very important difference between exposure of films to photons and electrons. Film exposed to photons exhibits a logarithmic response - that is, D varies as log(exposure). For electrons, this is not the case, but instead, the density is linear with exposure. This arises because individual grains require exposure to many photons to make them "developable", whereas a single electron will *COMPLETELY* expose many grains. For brevity, I will not explain how this changes the exposure characteristics, but it does.
A second consequence of this difference is that while change of development can change the threshold exposure for development of grains sensitised by light, grains exposed to electrons are either fully sensitized or are completely virgin, and there is *FAR* less scope to change the image characteristics by changing development of electron-exposed emulsions (though there is some, for Kodak SO163, for example).
This makes correct exposure much more critical for electron images, and makes them much more prone to overexposure. With light, and area with D of 4.0 has received 10,000 times more light than an area with D of 1.0. In an electron image the area with D of 4.0 has received 4 times more electrons than the area with D of 1.0. Do negatives really have D's above 4? Certainly they can when exposed to electrons.
There are other consequences. For example, the contrast (as we usually define it as the difference in density between different areas), which is independant of exposure on the linear portion of a film's response for light (as explained by Gary Gaugler), is, in the case of electrons, a linear function of the exposure. Underexposure leads to loss of contrast. This is why images of radiation-sensitive materials are taken at low magnification - it is the only way to maintain enough exposure of the emulsion to give acceptable contrast, without increasing the electron dose on the sample. Incidentally, the limit on information in such images is probably not the "grain size" of the emulsion, but the shot noise due to the finite number of electrons used to generate the image.
There is much more to this topic - and I have simplified what I have said.
Kristen is right on, especially with the safety issues. The completely dissolved agar should not look like it has sand in it when you swirl the flask. Be careful when swirling; if it is near the poiling point, it will foam up over the top before you can set it down. It can cause nasty burns like hot oil because it is thick.
Agar is a complex polysaccharide extracted from algae (several genera in the Rhodoophyceae). It melts somewhere around 96 degrees C and solidifies around 36 degrees C. I can't quite remember exact temperatures. Thus, to answer your solidification question, it will solidify when it gets to about body temperature. The length of time that takes after pouring will depend on how hot it was when it was poured, how thick your pour it, and how cool the Petri plate and room are.
Address at bottom if you have further questions.
On Tue, 17 Apr 2001, Kristen Lennon wrote:
} Date: Tue, 17 Apr 2001 11:48:59 -0500 } From: Kristen Lennon {kalen-at-iastate.edu} } To: Microscopy-at-sparc5.microscopy.com } Subject: Re: Ask-A-Microscopist:Agar } } ------------------------------------------------------------------------ } 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, } Your agar should be setting up almost immediately. I usually use a } concentration of 1-2% for my purposes. One imperative point is that the } agar needs to be completely dissolved. This may sound like a trivial } comment, but I know that when I first started working with agar I made the } mistake of not heating the solution hot enough for a long enough time and } wondered what was wrong. If you have access to an autoclave, autoclave it } for about 15 minutes. If not, heat it on a hot plate (preferably with a } stir bar, but you can swirl it periodically if you don't have a stir bar) } until the agar is completely dissolved and you have a nice, clear solution. } Be careful, because agar can suddenly start to boil and be up and over the } top of your flask before you realize it. } Let me know if you need any more help. } Good luck, } Kristen } } At 04:38 PM 4/16/01 -0500, you wrote: } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } } } Email: wonger-at-allover.com } } Name: Billone } } } } Organization: Murphy } } } } Education: 6-8th Grade Middle School } } } } Location: San Jose, California } } } } Question: How long should it take for agar to conjeal after I pour it into } } a petri dish? } } } } --------------------------------------------------------------------------- } } Kristen A. Lennon, Ph.D. } Department of Plant Pathology } 351 Bessey Hall } Iowa State University } Ames, IA 50011 } 515-294-8854 } kalen-at-iastate.edu } } }
Sara E. Miller, Ph. D. P. O. Box 3712 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-3265
I have been trying to locate a point source bulb for my Durst enlarger, It is GE bulb BHD it is a single contact bayonet type that is a 100 W , 20V bulb , which I have been told is an old microscope bulb that has been discontinued with no replacement noted. If anyone knows of or has a source where I can get a suitable replacement or have that bulb, please let me know.
Thanks,
Michael Pidgeon Keck School of Medicine at USC Dept. of Cell & Neurobiology
I have been trying to locate a point source bulb for my Durst enlarger, It is GE bulb BHD it is a single contact bayonet type that is a 100 W , 20V bulb , which I have been told is an old microscope bulb that has been discontinued with no replacement noted. If anyone knows of or has a source where I can get a suitable replacement or have that bulb, please let me know.
Thanks,
Michael Pidgeon Keck School of Medicine at USC Dept. of Cell & Neurobiology
Hello Everyone, I've written some software in Perl that may be of use in laboratory facilities. We use it to schedule and monitor usage of our TEM. It simulates the old sign up sheet on the wall idea, but on the web. It has a modest amount of security to control which users can sign up onto the schedule. It also keeps a searchable archive of your past schedules for history and billing information.
I've tried to make the installation as automated as possible. You'll need to have an account on a server where you can run cgi scripts, or your own linux/sun/irix/windows nt server. You'll also need to have Perl installed where you want the software to run. Please send me comments if you have any difficulties with setting up or running the software. I'm at version 1.2 which hopefully shouldn't have too many bugs.
The link for the software is at: http://wilfred.berkeley.edu/~gordon/www-sched I ask for a small fee for our lab for the software which is explained on the web page. Gordon Vrdoljak.
\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\/\ 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
Thanks to everyone who replied concerning my request for an EDS standardless quant. calibration standard for the TEM. NIST indeed do sell a mineral glass standard for just this purpose and I am looking into purchasing one of these. Thanks again for your help.
I have done some very nice preps using the Kleinschmit method. The procedure is a bit lengthy so please contact me off line and I would be more than happy to give it to you...It has some modifications but it works very nicely.
Sincerely, Maria
Maria Fazio-Zanakis AIM - TEM Laboratory Unilever Research, U.S. 45 River Road Edgewater, NJ 07020 1-201-840-2287 Maria.Fazio-Zanakis-at-unilever.com
Try Bulbtronics. Sorry, I don't know address, but I think they have a web site. They have all sorts of odd bulbs. Good luck.
On Tue, 17 Apr 2001, Michael Pidgeon wrote:
} Date: Tue, 17 Apr 2001 13:40:09 -0700 } From: Michael Pidgeon {pidgeon-at-hsc.usc.edu} } To: MS listserver {Microscopy-at-sparc5.microscopy.com} } Subject: Help locating replacment bulb } } ------------------------------------------------------------------------ } 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. } } I have been trying to locate a point source bulb for my Durst enlarger, } It is GE bulb BHD it is a single contact bayonet type that is a 100 W , } 20V bulb , which I have been told is an old microscope bulb that has } been discontinued with no replacement noted. If anyone knows of or has } a source where I can get a suitable replacement or have that bulb, } please let me know. } } Thanks, } } Michael Pidgeon } Keck School of Medicine at USC } Dept. of Cell & Neurobiology } } 323-442-1862 } } }
Sara E. Miller, Ph. D. P. O. Box 3712 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-3265
I will send you separately details of the method that I used on bacterial plasmids about 10-11 years ago (the method includes the preliminary stages as well as the technique). I can't send this to the list because the printed schedules need to be sent as attachments.
What I discovered was that it took several goes to get it right but the most important thing to avoid was 'surface active agents' which prevent a good film from spreading. Real 'double distilled water' is essential (not de-ionised or single distilled) and it should be stored in glass containers with no lubricants or plastic components (ground glass bottles or aluminium foil to prevent contact with plastic caps). The next was cleanliness - all glassware needs to be thoroughly soaked in a cleaning agent and then carefully rinsed with no use of detergents (I think some methods recommend chromic acid, but I found that a mixture of 2N (2M) nitric and 2N (1M) sulphuric acids were sufficient and a bit safer - NB obviously take care adding acids to water and do in a fume hood). Finally the 'hyperphase' containing the DNA should be mixed only a minute or 2 before use. Obviously to avoid contaminants it is based to keep a stock of chemicals just for this technique - but there aren't that many and they aren't particularly expensive.
Useful equipment would include a teflon 'Langmuir trough' to perform the spread (although I found that a square plastic dish worked well if thoroughly washed) and a rotary shadowing stage for the vacuum coater (although I just shadowed from two different angles at 90 deg of rotation - it takes longer because you have to return to air twice).
Once the technique was perfected I managed to get several dozen undergraduates to prepare DNA spreads of their plasmids, so it seemed like a fairly robust method where the important bit was the preparation before the DNA spread technique and a bit of practice. I found that in order for the students to get a representative number of plasmids it was best to photograph at about 10k and enlarge by 6x to 10x. I also found that the DNA was easiest to spot by increasing on-screen contrast in the microscope (smallest objective aperture and as low as 40kv electrons).
Malcolm
Malcolm Haswell e.m. unit University of Sunderland UK
Michael Jarnik 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 would need to prepare DNA for TEM using spreading/shadowing in } Cytochrome C films. My pilot experiments using just plasmid DNA and the } Lang & Mitani method (Biopolymers, 9, p.373, 1970) worked rather poorly } and I would like to hear from people with some experience in this } method. What are the critical points here? Purity of water/chemicals, } Cyt C concentration, time? Any hints would be appreciated. } } Thanks for help, } } -- } Michael Jarnik
Michael, The contact for Bulbtronics is 1-800-654-8542 or www.bulbtronics.com. They have a wide range of bulbs, including a 12 page catalog on just bulbs for microscopes.
I also have a file on a company called Bulb Direct. 1-800-772-5267 www.bulbdirect.com. They also have an extensive catalog.
I have no financial or other interest.............
Don Marshall
} From Microscopy-request-at-sparc5.microscopy.com Tue Apr 17 16:42:12 2001
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."
} } Hello All. } } I have been trying to locate a point source bulb for my Durst enlarger, } It is GE bulb BHD it is a single contact bayonet type that is a 100 W , } 20V bulb , which I have been told is an old microscope bulb that has } been discontinued with no replacement noted. If anyone knows of or has } a source where I can get a suitable replacement or have that bulb, } please let me know. } } Thanks, } } Michael Pidgeon } Keck School of Medicine at USC } Dept. of Cell & Neurobiology } } 323-442-1862 } }
They have an awesome selection of all sorts of bulbs at very good prices.
gary g.
At 01:38 PM 4/17/2001, you wrote:
} Hello All. } } I have been trying to locate a point source bulb for my Durst enlarger, } It is GE bulb BHD it is a single contact bayonet type that is a 100 W , } 20V bulb , which I have been told is an old microscope bulb that has } been discontinued with no replacement noted. If anyone knows of or has } a source where I can get a suitable replacement or have that bulb, } please let me know. } } Thanks, } } Michael Pidgeon } Keck School of Medicine at USC } Dept. of Cell & Neurobiology } } 323-442-1862
your argument about film density being linear with exposure for electrons came somewhat as a surprise. As you said, a single electron will *COMPLETELY* expose many grains. So, if the next electrons hits close to that area, some of the grains that would normally become developable are already completely exposed, thus the density of the film can not double and thus the response of the film is not linear.
There is a very simple test for the linearity of film if you have access to a field emission TEM with a biprism, or if you have a clean two beam case for high resolution. If you do a Fourier transform of just a cos-pattern, you should see two peaks in the Fourier transform. You can look this up in any handbook on Fourier transforms. If you do the same with a cos-pattern recorded on film, you will see plenty of higher order peaks in Fourier space. If you do this with a CCD camera, you will almost have to saturate the pixels to detect higher order terms. CCD cameras are, contrary to film, extremely linear.
If you have access to a film with a linear response, I would be eager to learn about it. Otherwise, for more details on my arguments, I would point to "Density Correction of Photographic Material ....", Ultramicroscopy, 55 (1994) 75-90. It gives examples of several films tested, the equations describing the behavior and tips how to linearize the data from film with software once the data are digitized.
With best regards,
Edgar Voelkl
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
--
________________ Dr. Edgar Voelkl Senior Development Staff Member ORNL Bldg 4515, MS 6064 1 Bethel Valley Road P.O. Box 2008 Oak Ridge, TN 37831-6064
Agar melts at 95 C and sets at ~40 C. If one melts agar (1-2%) directly on a hot plate, it can boil over when the temperature gets too high. However, if it is melted in a hot water bath, it will not boil over because the temperature will never get higher than 100 C.
How long does it take to set? It depends on the temperature of liquid agar when you pour it. If you pour 50 C agar, it will set immediately.
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
} } } "William F. Tivol" {wft03-at-health.state.ny.us} 04/17 8:48 AM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Email: wonger-at-allover.com Name: Billone
Organization: Murphy
Education: 6-8th Grade Middle School
Location: San Jose, California
Question: How long should it take for agar to conjeal after I pour it into a petri dish?
Dear Billone, Way back when I was pouring agar for immunodiffusion plates, I found that it congealed almost immediately. I had some trouble keeping it liquid to get a good smooth surface. Since that applies to the specific concentration of agar I used (I forget, but I think it was ~1%) and the temperature it was heated to (~40 C, I think), and since that was before global warming, YMMV. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
For electrons, this is not the case, but instead, the density is linear with exposure.
The only thing I will add to your consise explanation is that the density is linear with exposure only over a limited portion of the total dynamic range. The best films are linear for ODs from 0 to ~2, whereas the dynamic range is ~4 for these films. If one wants to determine the electron dose for the darker areas of the film, one must take a series of exposures to known doses of electrons and produce a curve of OD vs dose. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
} Does anyone know of service labs on the West coast for Amray } instruments? I would appreciate knowing if any exists.
A list of independent SEM service providers can be found at "www.jcnabity.com\service.htm".
Note that in some cases, these companies will service microscopes that are a significant distance from the company location. If a service area is not listed, you should contact the company to find out their service area.
If anyone has any additions or modifications that can be made to this list, please let me know.
Joe _________________________________________ Joe Nabity, Ph.D. JC Nabity Lithography Systems E-Beam Lithography using Commercial SEMs & STEMs PO Box 5354, Bozeman, MT 59717 USA Voice: (406) 587-0848 FAX: (406) 586-9514 E-mail: info-at-jcnabity.com Web: www.jcnabity.com
Sorry, Tony is correct about the linearity of film, at least after digitization. We have done this test many times, and provided the film is not grossly overexposed, the average exposure time and digitized counts are a straight line. (There is a small offset which you can correct for and is probably associated with our microdensitometer electronics rather than real.) Of course, if you push the film developing, you probably lost the linearity.
On Wed, 18 Apr 2001, Dr. Edgar Voelkl 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 Tony Garratt-Reed, } } your argument about film density being linear with exposure for } electrons came somewhat as a surprise. As you said, a single } electron will *COMPLETELY* expose many grains. So, if the next } electrons hits close to that area, some of the grains that would } normally become developable are already completely exposed, thus the } density of the film can not double and thus the response of the film } is not linear. } } There is a very simple test for the linearity of film if you have } access to a field emission TEM with a biprism, or if you have a clean } two beam case for high resolution. If you do a Fourier transform of } just a cos-pattern, you should see two peaks in the Fourier } transform. You can look this up in any handbook on Fourier } transforms. If you do the same with a cos-pattern recorded on film, } you will see plenty of higher order peaks in Fourier space. If you } do this with a CCD camera, you will almost have to saturate the } pixels to detect higher order terms. CCD cameras are, contrary to } film, extremely linear. } } If you have access to a film with a linear response, I would be eager } to learn about it. Otherwise, for more details on my arguments, I } would point to "Density Correction of Photographic Material ....", } Ultramicroscopy, 55 (1994) 75-90. It gives examples of several films } tested, the equations describing the behavior and tips how to } linearize the data from film with software once the data are } digitized. } } With best regards, } } Edgar Voelkl } } } } } } ------------------------------------------------------------------------ } } 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 } } -----------------------------------------------------------------------. } } } } } } Colleagues- } } } } I'm going out on a limb here, because it is not a particular area of } } expertise of mine, but I've not seen it in the discussion. Experts in } } microscopy of radiation sensitive materials are much more familiar with } } these issues (Linn Hobbs - can you help us here?) } } } } I'm surprised that no-one has pointed out a very important difference } } between exposure of films to photons and electrons. Film exposed to } } photons exhibits a logarithmic response - that is, D varies as } } log(exposure). For electrons, this is not the case, but instead, the } } density is linear with exposure. This arises because individual grains } } require exposure to many photons to make them "developable", whereas a } } single electron will *COMPLETELY* expose many grains. For brevity, I will } } not explain how this changes the exposure characteristics, but it does. } } } } A second consequence of this difference is that while change of development } } can change the threshold exposure for development of grains sensitised by } } light, grains exposed to electrons are either fully sensitized or are } } completely virgin, and there is *FAR* less scope to change the image } } characteristics by changing development of electron-exposed emulsions } } (though there is some, for Kodak SO163, for example). } } } } This makes correct exposure much more critical for electron images, and } } makes them much more prone to overexposure. With light, and area with D of } } 4.0 has received 10,000 times more light than an area with D of 1.0. In an } } electron image the area with D of 4.0 has received 4 times more electrons } } than the area with D of 1.0. Do negatives really have D's above 4? } } Certainly they can when exposed to electrons. } } } } There are other consequences. For example, the contrast (as we usually } } define it as the difference in density between different areas), which is } } independant of exposure on the linear portion of a film's response for } } light (as explained by Gary Gaugler), is, in the case of electrons, a } } linear function of the exposure. Underexposure leads to loss of contrast. } } This is why images of radiation-sensitive materials are taken at low } } magnification - it is the only way to maintain enough exposure of the } } emulsion to give acceptable contrast, without increasing the electron dose } } on the sample. Incidentally, the limit on information in such images is } } probably not the "grain size" of the emulsion, but the shot noise due to } } the finite number of electrons used to generate the image. } } } } There is much more to this topic - and I have simplified what I have said. } } } } Tony } } } } } } } } * * * * * * * * * * * * * * * * * * * * * * * * * * } } * Anthony J. Garratt-Reed M.A., D.Phil. } } * MIT, Room 13-1027 } } * 77 Massachusetts Avenue } } * Cambridge, MA 02139-4307 } } * USA } } * Phone: (617) 253-4622 } } * Fax: (617) 258-6478 } } * } } -- } } } ________________ } Dr. Edgar Voelkl } Senior Development Staff Member } ORNL } Bldg 4515, MS 6064 } 1 Bethel Valley Road } P.O. Box 2008 } Oak Ridge, TN 37831-6064 } } Tel.: (865) 574-8181 } Fax: (865) 574-4913 } email: vog-at-ornl.gov } }
------------------------------------------------------- Laurence Marks Department of Materials Science and Engineering & Center for Transportation Nanotechnology Northwestern University Tel: (847) 491-3996 Fax: (847) 491-7820 mailto:ldm-at-risc4.numis.nwu.edu http://www.numis.nwu.edu http://www.ctn.northwestern.edu ------------------------------------------------------- The Other Nanotubes http://focus.aps.org/open/st12.html Boron Nitride Nanotubes http://pubs.acs.org/cen/topstory/7912/7912notw1.html
Workshop May 17-19 2001 "New approaches to the Phase Problem" http://xraysweb.lbl.gov/esg/phasing/index.html
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Sorry too :)
however I feel quite confident that I can stand behind my experimental results. And I can assure you that film developing was not pushed. In addition, when digitizing film, you get a second non-linear function involved. Right? You can check this by using a Kodak photographic step tablet no.3. They are available off the shelf (at least used to be) and are calibrated. Density range is 21 steps from 0.05 to 3.05. But, maybe your microdensitometer is already corrected for the transmittivity curve (Pixel value P = a 10**S, where a corresponds to the illumination intensity and S is the density of the film) ?
Maybe, we are just talking degrees of linearity here? So, whatever you have may be sufficient in some cases, but no in others?
Edgar
} } Sorry, } Tony is correct about the linearity of film, at least after } digitization. We have done this test many times, and provided the film } is not grossly overexposed, the average exposure time and digitized } counts are a straight line. (There is a small offset which you can } correct for and is probably associated with our microdensitometer } electronics rather than real.) } Of course, if you push the film developing, you probably lost } the linearity. } } } On Wed, 18 Apr 2001, Dr. Edgar Voelkl 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 Tony Garratt-Reed, } } } } your argument about film density being linear with exposure for } } electrons came somewhat as a surprise. As you said, a single } } electron will *COMPLETELY* expose many grains. So, if the next } } electrons hits close to that area, some of the grains that would } } normally become developable are already completely exposed, thus the } } density of the film can not double and thus the response of the film } } is not linear. } } } } There is a very simple test for the linearity of film if you have } } access to a field emission TEM with a biprism, or if you have a clean } } two beam case for high resolution. If you do a Fourier transform of } } just a cos-pattern, you should see two peaks in the Fourier } } transform. You can look this up in any handbook on Fourier } } transforms. If you do the same with a cos-pattern recorded on film, } } you will see plenty of higher order peaks in Fourier space. If you } } do this with a CCD camera, you will almost have to saturate the } } pixels to detect higher order terms. CCD cameras are, contrary to } } film, extremely linear. } } } } If you have access to a film with a linear response, I would be eager } } to learn about it. Otherwise, for more details on my arguments, I } } would point to "Density Correction of Photographic Material ....", } } Ultramicroscopy, 55 (1994) 75-90. It gives examples of several films } } tested, the equations describing the behavior and tips how to } } linearize the data from film with software once the data are } } digitized. } } } } With best regards, } } } } Edgar Voelkl } } } } } } } } } } } ------------------------------------------------------------------------ } } } 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 } } } -----------------------------------------------------------------------. } } } } } } } } } Colleagues- } } } } } } I'm going out on a limb here, because it is not a particular area of } } } expertise of mine, but I've not seen it in the discussion. Experts in } } } microscopy of radiation sensitive materials are much more familiar with } } } these issues (Linn Hobbs - can you help us here?) } } } } } } I'm surprised that no-one has pointed out a very important difference } } } between exposure of films to photons and electrons. Film exposed to } } } photons exhibits a logarithmic response - that is, D varies as } } } log(exposure). For electrons, this is not the case, but instead, the } } } density is linear with exposure. This arises because individual grains } } } require exposure to many photons to make them "developable", whereas a } } } single electron will *COMPLETELY* expose many grains. For brevity, I will } } } not explain how this changes the exposure characteristics, but it does. } } } } } } A second consequence of this difference is that while change of development } } } can change the threshold exposure for development of grains sensitised by } } } light, grains exposed to electrons are either fully sensitized or are } } } completely virgin, and there is *FAR* less scope to change the image } } } characteristics by changing development of electron-exposed emulsions } } } (though there is some, for Kodak SO163, for example). } } } } } } This makes correct exposure much more critical for electron images, and } } } makes them much more prone to overexposure. With light, and area with D of } } } 4.0 has received 10,000 times more light than an area with D of 1.0. In an } } } electron image the area with D of 4.0 has received 4 times more electrons } } } than the area with D of 1.0. Do negatives really have D's above 4? } } } Certainly they can when exposed to electrons. } } } } } } There are other consequences. For example, the contrast (as we usually } } } define it as the difference in density between different areas), which is } } } independant of exposure on the linear portion of a film's response for } } } light (as explained by Gary Gaugler), is, in the case of electrons, a } } } linear function of the exposure. Underexposure leads to loss of contrast. } } } This is why images of radiation-sensitive materials are taken at low } } } magnification - it is the only way to maintain enough exposure of the } } } emulsion to give acceptable contrast, without increasing the electron dose } } } on the sample. Incidentally, the limit on information in such images is } } } probably not the "grain size" of the emulsion, but the shot noise due to } } } the finite number of electrons used to generate the image. } } } } } } There is much more to this topic - and I have simplified what I have said. } } } } } } Tony } } } } } } } } } } } } * * * * * * * * * * * * * * * * * * * * * * * * * * } } } * Anthony J. Garratt-Reed M.A., D.Phil. } } } * MIT, Room 13-1027 } } } * 77 Massachusetts Avenue } } } * Cambridge, MA 02139-4307 } } } * USA } } } * Phone: (617) 253-4622 } } } * Fax: (617) 258-6478 } } } * } } } } -- } } } } } } ________________ } } Dr. Edgar Voelkl } } Senior Development Staff Member } } ORNL } } Bldg 4515, MS 6064 } } 1 Bethel Valley Road } } P.O. Box 2008 } } Oak Ridge, TN 37831-6064 } } } } Tel.: (865) 574-8181 } } Fax: (865) 574-4913 } } email: vog-at-ornl.gov } } } } } } ------------------------------------------------------- } Laurence Marks } Department of Materials Science and Engineering & } Center for Transportation Nanotechnology } Northwestern University } Tel: (847) 491-3996 Fax: (847) 491-7820 } mailto:ldm-at-risc4.numis.nwu.edu } http://www.numis.nwu.edu http://www.ctn.northwestern.edu } ------------------------------------------------------- } The Other Nanotubes http://focus.aps.org/open/st12.html } Boron Nitride Nanotubes http://pubs.acs.org/cen/topstory/7912/7912notw1.html } } Workshop May 17-19 2001 "New approaches to the Phase Problem" } http://xraysweb.lbl.gov/esg/phasing/index.html
--
________________ Dr. Edgar Voelkl Senior Development Staff Member ORNL Bldg 4515, MS 6064 1 Bethel Valley Road P.O. Box 2008 Oak Ridge, TN 37831-6064
Very often it seems that film is described as non-linear, direct CCD collectors are linear, so bye-bye film. Since we did not then (and still do not have) a good CCD detector on our UHV-HREM, we investigated this in detail some years ago to get quantitative TED data from surfaces. While there are of course logarithmic terms involved, with a good drum scanner the electronics to handle this works well. If anyone wants we have calibration curves sitting on the wall over our scanner and I can send them. So long as you maintain the microdensitometer, performing occaisional calibrations, everything is fine. (A microdensitometer without reasonable TLC - GIGO.)
Both CCD camera's on the microscope and scanners have point-spread functions, and these can be important. For instance, we have a rotating drum scanner so the PSF is different along the rotation direction versus at right angles to it. I do not know what the PSF's for modern scanners are like - hence the question I raised a little time ago (but nobody had more information). You also have to have adequate sampling and similar "stuff" under control otherwise aliasing kills you.
A 2kx2k CCD is certainly competitive with film, depending upon the application. Scratches and damage from high intensities are problems for CCD's, but having an image immediately available so you can look at the power-spectrum is an advantage. However, I think the price of the TEM units has to come down before film and $2k (or even $20k) scanners become obsolete.
On Wed, 18 Apr 2001, Dr. Edgar Voelkl 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 } } -----------------------------------------------------------------------. } } Sorry too :) } } however I feel quite confident that I can stand behind my } experimental results. And I can assure you that film developing was } not pushed. In addition, when digitizing film, you get a second } non-linear function involved. Right? You can check this by using a } Kodak photographic step tablet no.3. They are available off the } shelf (at least used to be) and are calibrated. Density range is 21 } steps from 0.05 to 3.05. But, maybe your microdensitometer is } already corrected for the transmittivity curve (Pixel value P = a } 10**S, where a corresponds to the illumination intensity and S is the } density of the film) ? } } Maybe, we are just talking degrees of linearity here? So, whatever } you have may be sufficient in some cases, but no in others? } } Edgar } } } } } } } Sorry, } } Tony is correct about the linearity of film, at least after } } digitization. We have done this test many times, and provided the film } } is not grossly overexposed, the average exposure time and digitized } } counts are a straight line. (There is a small offset which you can } } correct for and is probably associated with our microdensitometer } } electronics rather than real.) } } Of course, if you push the film developing, you probably lost } } the linearity. } } } } } } On Wed, 18 Apr 2001, Dr. Edgar Voelkl 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 Tony Garratt-Reed, } } } } } } your argument about film density being linear with exposure for } } } electrons came somewhat as a surprise. As you said, a single } } } electron will *COMPLETELY* expose many grains. So, if the next } } } electrons hits close to that area, some of the grains that would } } } normally become developable are already completely exposed, thus the } } } density of the film can not double and thus the response of the film } } } is not linear. } } } } } } There is a very simple test for the linearity of film if you have } } } access to a field emission TEM with a biprism, or if you have a clean } } } two beam case for high resolution. If you do a Fourier transform of } } } just a cos-pattern, you should see two peaks in the Fourier } } } transform. You can look this up in any handbook on Fourier } } } transforms. If you do the same with a cos-pattern recorded on film, } } } you will see plenty of higher order peaks in Fourier space. If you } } } do this with a CCD camera, you will almost have to saturate the } } } pixels to detect higher order terms. CCD cameras are, contrary to } } } film, extremely linear. } } } } } } If you have access to a film with a linear response, I would be eager } } } to learn about it. Otherwise, for more details on my arguments, I } } } would point to "Density Correction of Photographic Material ....", } } } Ultramicroscopy, 55 (1994) 75-90. It gives examples of several films } } } tested, the equations describing the behavior and tips how to } } } linearize the data from film with software once the data are } } } digitized. } } } } } } With best regards, } } } } } } Edgar Voelkl } } } } } } } } } } } } } } } } ------------------------------------------------------------------------ } } } } 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 } } } } -----------------------------------------------------------------------. } } } } } } } } } } } } Colleagues- } } } } } } } } I'm going out on a limb here, because it is not a particular area of } } } } expertise of mine, but I've not seen it in the discussion. Experts in } } } } microscopy of radiation sensitive materials are much more familiar with } } } } these issues (Linn Hobbs - can you help us here?) } } } } } } } } I'm surprised that no-one has pointed out a very important difference } } } } between exposure of films to photons and electrons. Film exposed to } } } } photons exhibits a logarithmic response - that is, D varies as } } } } log(exposure). For electrons, this is not the case, but instead, the } } } } density is linear with exposure. This arises because individual grains } } } } require exposure to many photons to make them "developable", whereas a } } } } single electron will *COMPLETELY* expose many grains. For brevity, I will } } } } not explain how this changes the exposure characteristics, but it does. } } } } } } } } A second consequence of this difference is that while change of development } } } } can change the threshold exposure for development of grains sensitised by } } } } light, grains exposed to electrons are either fully sensitized or are } } } } completely virgin, and there is *FAR* less scope to change the image } } } } characteristics by changing development of electron-exposed emulsions } } } } (though there is some, for Kodak SO163, for example). } } } } } } } } This makes correct exposure much more critical for electron images, and } } } } makes them much more prone to overexposure. With light, and area with D of } } } } 4.0 has received 10,000 times more light than an area with D of 1.0. In an } } } } electron image the area with D of 4.0 has received 4 times more electrons } } } } than the area with D of 1.0. Do negatives really have D's above 4? } } } } Certainly they can when exposed to electrons. } } } } } } } } There are other consequences. For example, the contrast (as we usually } } } } define it as the difference in density between different areas), which is } } } } independant of exposure on the linear portion of a film's response for } } } } light (as explained by Gary Gaugler), is, in the case of electrons, a } } } } linear function of the exposure. Underexposure leads to loss of contrast. } } } } This is why images of radiation-sensitive materials are taken at low } } } } magnification - it is the only way to maintain enough exposure of the } } } } emulsion to give acceptable contrast, without increasing the electron dose } } } } on the sample. Incidentally, the limit on information in such images is } } } } probably not the "grain size" of the emulsion, but the shot noise due to } } } } the finite number of electrons used to generate the image. } } } } } } } } There is much more to this topic - and I have simplified what I have said. } } } } } } } } Tony } } } } } } } } } } } } } } } } * * * * * * * * * * * * * * * * * * * * * * * * * * } } } } * Anthony J. Garratt-Reed M.A., D.Phil. } } } } * MIT, Room 13-1027 } } } } * 77 Massachusetts Avenue } } } } * Cambridge, MA 02139-4307 } } } } * USA } } } } * Phone: (617) 253-4622 } } } } * Fax: (617) 258-6478 } } } } * } } } } } } -- } } } } } } } } } ________________ } } } Dr. Edgar Voelkl } } } Senior Development Staff Member } } } ORNL } } } Bldg 4515, MS 6064 } } } 1 Bethel Valley Road } } } P.O. Box 2008 } } } Oak Ridge, TN 37831-6064 } } } } } } Tel.: (865) 574-8181 } } } Fax: (865) 574-4913 } } } email: vog-at-ornl.gov } } } } } } } } } } ------------------------------------------------------- } } Laurence Marks } } Department of Materials Science and Engineering & } } Center for Transportation Nanotechnology } } Northwestern University } } Tel: (847) 491-3996 Fax: (847) 491-7820 } } mailto:ldm-at-risc4.numis.nwu.edu } } http://www.numis.nwu.edu http://www.ctn.northwestern.edu } } ------------------------------------------------------- } } The Other Nanotubes http://focus.aps.org/open/st12.html } } Boron Nitride Nanotubes http://pubs.acs.org/cen/topstory/7912/7912notw1.html } } } } Workshop May 17-19 2001 "New approaches to the Phase Problem" } } http://xraysweb.lbl.gov/esg/phasing/index.html } } -- } } } ________________ } Dr. Edgar Voelkl } Senior Development Staff Member } ORNL } Bldg 4515, MS 6064 } 1 Bethel Valley Road } P.O. Box 2008 } Oak Ridge, TN 37831-6064 } } Tel.: (865) 574-8181 } Fax: (865) 574-4913 } email: vog-at-ornl.gov } }
------------------------------------------------------- Laurence Marks Department of Materials Science and Engineering & Center for Transportation Nanotechnology Northwestern University Tel: (847) 491-3996 Fax: (847) 491-7820 mailto:ldm-at-risc4.numis.nwu.edu http://www.numis.nwu.edu http://www.ctn.northwestern.edu ------------------------------------------------------- The Other Nanotubes http://focus.aps.org/open/st12.html Boron Nitride Nanotubes http://pubs.acs.org/cen/topstory/7912/7912notw1.html
Workshop May 17-19 2001 "New approaches to the Phase Problem" http://xraysweb.lbl.gov/esg/phasing/index.html
Ed Voelkl is a good microscopist whom I like and respect but he is wrong. Of course, all detectors saturate if the signal is large enough. But the point is how does it behave in the useful detection range. When film is exposed to electrons the density increases linearly with dose - until saturation effects begin. By contrast when exposed to light, film is not linear anywhere.
If you want to get this right, please read:
The response of photographic emulsions to electrons by R C Valentine in Advances in Optical and Electron Microscopy, volume 1 Edited by R Barer and V E Cosslett Academic Press
This is one of those definitive and good articles - which is also clear and easy to read.
Alwyn Eades -- .......... Alwyn Eades Department of Materials Science and Engineering Lehigh University 5 East Packer Avenue Bethlehem Pennsylvania 18015-3195 Phone 610 758 4231 Fax 610 758 4244 jae5-at-lehigh.edu
Hey, I am enjoying this discussion. Any chance to get a lunchtime debate going at the M&M meeting like we had a few years ago?
-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: Alwyn Eades [mailto:jae5-at-lehigh.edu] } Sent: Wednesday, April 18, 2001 2:55 PM } To: EMNET } Subject: Exposure of film to electrons. } } } -------------------------------------------------------------- } ---------- } 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/MicroscopyLists } erver/FAQ.html } } } } -------------------------------------------------------------- } ---------. } } } Ed Voelkl is a good microscopist whom I like and respect but he is } wrong. Of course, all detectors saturate if the signal is large } enough. But the point is how does it behave in the useful detection } range. When film is exposed to electrons the density } increases linearly } with dose - until saturation effects begin. By contrast when exposed } to light, film is not linear anywhere. } } If you want to get this right, please read: } } The response of photographic emulsions to electrons } by R C Valentine } in Advances in Optical and Electron Microscopy, volume 1 } Edited by R Barer and V E Cosslett } Academic Press } } This is one of those definitive and good articles - which is } also clear } and easy to read. } } Alwyn Eades } -- } .......... } Alwyn Eades } Department of Materials Science and Engineering } Lehigh University } 5 East Packer Avenue } Bethlehem } Pennsylvania 18015-3195 } Phone 610 758 4231 } Fax 610 758 4244 } jae5-at-lehigh.edu }
Mary McCann's email address is: mccanns-at-tiac.net. I'm sorry for any inconvenience to those people who have tried, unsuccessfully, to reach her!
If anyone is interested in attending the meeting--you can email me directly and I can send you a newsletter or fax you the information re: program and registration.
Please include a complete mailing address and fax number.
Thanks! Peggy Sherwood Corresponding Secretary, NESM -- Peggy Sherwood Lab Associate, Photopathology Wellman Laboratories of Photomedicine (W224) Massachusetts General Hospital 50 Blossom Street Boston, MA 02114 617-724-4839 (voice mail) 617-726-6983 (lab) 617-726-3192 (fax) sherwood-at-helix.mgh.harvard.edu
H. Frieser and E. Klein, Z. Angew. Phys. 10 (1958) 337.
H. Frieser, E. Klein and E. Zeitler, Z. Angew. Phys. 11 (1959) 190.
E. Zeitler, Ultramicroscopy 46 (1992) 405.
E. Voelkl, F. Lenz, Q. Fu and H. Lichte, "Density correction of photographic material for further image processing ....", UM 55 (1994) 75-89.
against:
R C Valentine, in Advances in Optical and Electron Microscopy, volume 1 Edited by R Barer and V E Cosslett, Academic Press
that's 4:1 :)
Edgar
P.S.: So far, the argument remains: a single electron "A" will completely expose many grains. So, if the next electron "B" hits close to the same area, some of the grains are already completely exposed through electron "A", thus less grains become fully developed with electron "B". Therefore, the density of the film can not double and thus the response of the film is non-linear.
At 2:55 PM -0400 4/18/01, Alwyn Eades wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
--
________________ Dr. Edgar Voelkl Senior Development Staff Member ORNL Bldg 4515, MS 6064 1 Bethel Valley Road P.O. Box 2008 Oak Ridge, TN 37831-6064
Since this seems to be a lively thread, I would like to make the following suggestion:
Most of us have an electron microscope, and I would like to suggest the following experiment to test if your film is linear or not. It is a simple, two-step test.
1) Take a high resolution image of some periodic structure such that only two beams contribute to the image, preferably with identical intensity. This will provide a cos- type imprint on your film (actually a P+cos(...) type imprint with P} 1).
2) Develop the film and look through it at a small light source, e.g., a halogen light. If necessary, tilt the film a little to decrease fringe spacing for better separation of the diffraction orders.
If you have a true linear film, no higher diffraction orders will be visible. The two diffracted beams will be quite colorful like a rainbow. If you see more than two diffracted beams, your film has a non-linear response (all of those I have ever seen do (disclaimer: the last time I have dealt with film was 1992)).
BTW,
I like Scotts suggestion of a lunchtime debate going at the M&M meeting. Monday is out with me, but any other day is fine.
Edgar
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
--
________________ Dr. Edgar Voelkl Senior Development Staff Member ORNL Bldg 4515, MS 6064 1 Bethel Valley Road P.O. Box 2008 Oak Ridge, TN 37831-6064
Amray has a list of third party repair outfits. I got it from Amray. I can fax it to you or call Amray and get it direct.
gary g.
At 11:20 AM 4/17/2001, you wrote:
} Does anyone know of service labs on the West coast for Amray } instruments? I would appreciate knowing if any exists. } Thank you. } } Franklin Bailey } University of Idaho } Moscow, ID 83844-2204 } jfb-at-uidaho.edu
Quick and dirty way to check the linearity of the whole acquisition process (film + scanner) is to take multiple exposures on a film by gradually exposing its area. I've done this by using the selected area aperture. First position the aperture so that only a small stripe of the film to be exposed. Make an exposure. Then move the aperture to open a larger area (next stripe). Expose ... Use a relatively low dose so that after 20 or 30 stripes the film to be exposed to twice the dose you use in your experiments. After scanning the film make a "line projection". Plotting the density vs. stripe number (dose) will show you the linearity region of your acquisition system (film + scanner). A quick way to check the Modulation Transfer Function (MTF) of the acquisition system is to take an exposure on the film without specimen (beam only). Theoretically the Fourier transform of the scan should show a constant amplitude over all spatial frequencies (Shot noise = white noise). Average rotationally the amplitude of the Fourier transform - this is the MTF.
Does anyone know of a digital imaging system for scanning EM? I know that CCD imaging systems for TEMs have been discussed in the group before, but I don't recall whether SEMs have been discussed. Perhaps someone could tell me if this subject has been archived, or perhaps someone has a summary. Vendors should also feel free to respond directly to me off-list.
Someone has asked me for information on a digital image acquisition system for SEM (they currently have a screen with a Polaroid camera attachment). If anyone has recommendations for such a digital acquisition system, please contact me with the details.
Thanks very much in advance.
Bob Chiovetti GTI Microsystems, Inc. Leica Exclusive Regional Dealer Southwestern United States rchiovetti-at-aol.com
Just let you know that there will be two short courses on food microscopy in Minneapolis on May 13 as part of the Food Structure & Functionality Symposium.
Course 1 is aimed at researchers interested in specific localisation techniques as a tool for understanding structure-function relationships in foods. Course 2 gives a grounding in light microscopy technques (optical contrast, staining) used for the identification of food materials and contaminants (glass, fibres, plastics etc) and is aimed at quality control/research & development personnel.
Details can be found at: http://www.aocs.org/meetings/am2001/foodstr.htm
Both courses offer practical tuition by internationally recognised speakers.
Let me know if you are interested, or you can book online at: https://www.aocs.org/meetings/am2001/regshort.htm
Regards Mark
Mark Auty Dairy Products Research Centre Moorepark Fermoy Co. Cork Ireland
tel +353 25 42447 fax +353 25 42340 mauty-at-moorepark.teagasc.ie
Just let you know that there will be two short courses on food microscopy in Minneapolis on May 13 as part of the Food Structure & Functionality Symposium.
Course 1 is aimed at researchers interested in specific localisation techniques as a tool for understanding structure-function relationships in foods. Course 2 gives a grounding in light microscopy technques (optical contrast, staining) used for the identification of food materials and contaminants (glass, fibres, plastics etc) and is aimed at quality control/research & development personnel.
Details can be found at: http://www.aocs.org/meetings/am2001/foodstr.htm
Both courses offer practical tuition by internationally recognised speakers.
Let me know if you are interested, or you can book online at: https://www.aocs.org/meetings/am2001/regshort.htm
Regards Mark
Mark Auty Dairy Products Research Centre Moorepark Fermoy Co. Cork Ireland
tel +353 25 42447 fax +353 25 42340 mauty-at-moorepark.teagasc.ie
Hi, I would like to get in contact with anyone currently using a JEOL-220A SEM. I particularly want to know if it is possible to add a digital image acquisition system to this SEM. Our lab has a chance to acquire a 220A which has less than 200 hours of use and was under service contract before going into storage 4 years ago. Any and all information would be appreciated. Thanks.
Tom Bargar EM Lab U. Neb. Med. Ctr. phone (402)-559-7347 tbargar-at-unmc.edu
The Biomaterials Science Group Department of Oral and Dental Science University of Bristol in collaboration with Glaxo SmithKline
Postdoctoral Research Assistant
in the Biomaterials Science Group on the project Interaction Mechanisms of Polymers at Interfaces of Mineralised Tissues
The research area involves the study of the physical and chemical properties and interaction mechanisms of different polymers at interfaces of mineralised tissues. You will have recently been awarded a PhD in an appropriate field and will ideally have experience in scanning probe microscopy (AFM) of biological materials and other analytical techniques and an interest in medical research. You will work in Dr. Jandts group and interact with scientists at Glaxo Smith Kline. The University of Bristol is one of the leading research universities in the UK and provides an outstanding scientific training environment to enhance your qualification. The group is involved in exciting, interdisciplinary projects and maintains appropriate state of the art instrumentation. There exist opportunities for additional interactions with clinical scientists and other centres at the university. We are looking for a dynamic and exceptionally well-qualified postdoctoral researcher who can interact effectively in an international and interdisciplinary team. The appointment will be on a Research Assistant 1A scale with a salary range of # 16775 to # 20465. This is a full time appointment and initially for one year. Applicants should include a short CV, stating research experience and interests, publication list and addresses of two referees. The review of applications will start 24 May 2001 and will continue until the post has been filled. Informal inquiries can be directed by email to Dr. K. D. Jandt (K.Jandt-at-bris.ac.uk), Senior Lecturer in Biomaterials, University of Bristol
Formal applications quoting the reference number 7401 should be directed to
The University of Bristol Recruitment Office Bristol, BS8 1TH United Kingdom
----------------------------------------------------------------- Dr. rer. nat. Klaus D. Jandt Senior Lecturer in Dental Materials Science and Biomaterials University of Bristol, Department of Oral and Dental Science Lower Maudlin Street, Bristol, BS1 2LY, UK Phone: +44-117-9284418, Fax: ++44-117-9284780 Internet: K.Jandt-at-bris.ac.uk WWW: http://www.dent.bris.ac.uk/Biomaterials/kdj.htm "We make Biomaterials Science work!"
I have managed to get a Varian 842 ionization gauge controller and am working on the tube itself, leaving only one component - the cable(s) - remaining. Varian wants $190 for this cable, which seems a little steep to me.
} From the connections on the controller box it looks like there is a four wire cable with an octal plug going that would go to some connector on the bottom of the tube and a separate coax connection for the collector. Are the connectors that would attach to the bottom and top of the tube readily available components and, if so, where could I buy them? Is there some place where I could simply buy the entire cable system at a better price than what Varian is asking? What is so special about five wires and a couple of connectors that warrants that kind of price?
I would appreciate information from anyone regarding the connection of ion gauge tubes to their controllers, Varian or not.
ASSISTANT DIRECTOR (TECHNICAL) Acadia Centre for Microstructural Analysis (ACMA) Competition #01-17
The Acadia Centre for Microstructural Analysis (ACMA) has recently been established to support research initiatives within the Faculty of Pure & Applied Science and provide technical services to the regional R&D community. We are seeking an individual to manage and maintain the microanalytical facility consisting of scanning and transmission electron microscopes, scanning probe microscope, FTIR spectrometer, epi-fluorescence and confocal microscopes along with associated specimen preparation equipment.
Responsibilities: management of day to day operations including training and general supervision of users, technical support and general maintenance of research equipment, technical service and consulting activities, marketing of technical services.
Qualifications: A minimum of a Masters Degree in natural sciences/engineering with courses in computer programming and electronics or an equivalent combination of education and related work experience; experience in instrumentation; some expertise with one or more of TEM, SEM, FTIR, SPM or confocal microscopy; strong communication, writing and interpersonal skills. An ability to provide general electronics and instrumentation support to the Faculty would be considered an asset.
The Assistant Director will be a full-time employee of Acadia University and the initial appointment shall be for one year, with the possibility of renewal for up to five years.
Salary Range: $39,615 to $51,645 per annum, depending on qualifications
Closing Date for Applications: June 1, 2001
Please send your resume including three letters of references to: Marian Reid, Personnel Officer Acadia University, Wolfville NS B0P 1X0 E-mail: marian.reid-at-acadiau.ca Fax: 902-585-1075
We thank all applicants in advance but advise that only those selected for an interview will be contacted. Acadia University reserves the right not to fill this position.
In accordance with Canadian immigration requirements, this advertisement is directed to Canadian citizens and permanent residents.
An equal opportunity employer, Acadia welcomes applications from qualified women and men, including African Nova Scotians, First Nations peoples, persons with disabilities, and racially visible people. Such individuals are encouraged to self-identify.
Dr. Craig Bennett Associate Professor and Acting Head Department of Physics, Acadia University and Director, Acadia Centre for Microstructural Analysis Wolfville, Nova Scotia, Canada B0P 1X0 tel. 902-585-1150 fax. 902-585-1816
Hi all! I have a quick question. I am working with purified nuclear matrix samples and need tips for the fixation / embedding process. First of all, the samples are so small, they are nearly invisible, and secondly, they are incredibly fragile, so great care must be taken during this whole process. To complicate matters, we are doing pre-embedding immunolabeling. The problem comes in after incubation in secondary (gold) antibody. We have to spin down the sample after each rinse or incubation. My concern is about the gold particles........At what RPM will they sediment? Has anyone ever encountered this dileama before? Likewise, does anyone have any tips for fixation and embedding of this kind of sample? Question # 2: Is it ok to go straight from Prop Ox to 100% resin (spurr's)? The matrix is mostly protein, so it shouldn't take long at all to infiltrate...but I am unsure about how to go about infiltrating. In the viscous resin, we won't be able to spin down the matrix, that's where the problem arises.
Any help would be greatly appreciated!
Thanks in advance, Karli Karli Fitzelle MCI Center OARDC / OSU Wooster, Ohio 44691 330-263-3828
I have a user in our EM Center who would like to have a procedure for determining the thickness of their ultrathin sections, 60-100 nm. They are using the normal color criterion of "silver" sections in the boat to select their sections, but would like to be more precise. There is a potential for doing morphometry and comparisons of particle counts between non-serial sections and sections from different specimens.
They have heard of a technique that uses small particles applied to both surfaces of the section and using tilt and geometry of the TEM stage to determine section thickness. Any details of this technique would be appreciated.
All suggestions are welcome.
John Colorado State University john.chandler-at-colostate.edu
Sample holders for JEOL 100CX-2000EXII model TEMs for sale:
Gatan hot stage sample holders, air/water cooled. Both furnaces in working order; holders include one hexnut lockring AND thermocouple controller (sorry, no power wires--lost!): --#1 in good condition, $12,000 (Gatan Power supply model #580-0300) --#2 lightly stripped hex nut assembly in furnace (doesn't tighten all the way but sample will remain in place fairly well), mA display on power supply (Gatan power supply model #628-0500) a bit jumpy but thermocouple reads temperature fine, $7000
Extra washers (Gatan part #628-0223), and hex nut tool (Gatan part #608-0005) NOT included with the above holders.
Please contact Terry K. Baker for further inquiries and negotiations: Phone 508 893 9560 baker-at-catalytic-materials.com Catalytic Materials 1750 Washington St. West Holliston Professional Park, Suite C2 Holliston, MA 01746
I would try to immobilize the stuff on some substrate that could then be carried like a piece of tissue. I am not sure what that would be in this case. Possibly a Nuclepore filter???
At 03:27 PM 4/19/2001 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Greg Erdos Assistant Director Biotechnology Program Ph. 352-392-1295 University of Florida Fax 352-846-0251 PO Box 118525 Gainesville, FL 32611 http://www.biotech.ufl.edu/~emcl
I have been asked by one of our collegues hereas to wether there is a selective stain for viewing xylem tissue that has been embedded in resin. These are 1-7um sections viewed with a light microscope
Somewhere in the deep dark recesses (which are not accessible on a Friday afternoon or Monday morning ) I seem to remember there is one; or indeed, a publication that lists many of these such permeations.
Any help appreciated Cheers
Raymond Bennett Keith Williamson EM Unit Hort+Research Palmerston North NEW ZEALAND
_________________________________________________________________ The contents of this e-mail are privileged and/or confidential to the named recipient and are not to be used by any other person and/or organisation. If you have received this e-mail in error, please notify the sender and delete all material pertaining to this e-mail. _________________________________________________________________
} I have managed to get a Varian 842 ionization gauge controller and am } working on the tube itself, leaving only one component - the cable(s) - } remaining. Varian wants $190 for this cable, which seems a little steep to } me. } } } From the connections on the controller box it looks like there is a four } wire cable with an octal plug going that would go to some connector on the } bottom of the tube and a separate coax connection for the collector. Are the } connectors that would attach to the bottom and top of the tube readily } available components and, if so, where could I buy them? Is there some place } where I could simply buy the entire cable system at a better price than what } Varian is asking? What is so special about five wires and a couple of } connectors that warrants that kind of price? } } I would appreciate information from anyone regarding the connection of ion } gauge tubes to their controllers, Varian or not. } } Bruce Girrell
I got a similar cable from Duniway Stockroom. They were much cheaper than Varian, and the cable has worked great.
Tom
-- Thomas Mullarkey Murray email:tm8a-at-virginia.edu Thornton Hall - MSE phone:(804)982-5659 University of Virginia Fax: (804)982-5660 Charlottesville, VA 22903
Earl ----- Original Message ----- } From: "Bruce Girrell" {bigirrell-at-microlinetc.com} To: {Microscopy-at-sparc5.microscopy.com} Sent: Thursday, April 19, 2001 9:26 AM
Question: I am learning to do SEM now and have been doing immunolocalization (histogold) for light microscopy. I am now interested in applying this histogold immunolocalization in EM work. It appears that most immunolocalization I've seen is in TEM. I am working in SEM. Is there any advantage of one over the other besides the obvious difference that TEM looks inside the tissue and SEM looks at the surface? How come there aren't as many SEM localizations out there (that I've seen)? Thank you for your time, Karen
} Subject: EDXA, Need help with detector geometry } } } } Sirs or Madams, } } } } I am running a JEOL CX II with a Kevex mod. 3200-0018 detector/ Kevex Delta } } Class Anlyzer. } } } } I am having difficulty locating the geometric variables unique to this } } mating of scope and detector. Kevex was unable to supply the data. These } } geometric variables are used by the analyzer software (Quantex) in modeling } } and subtracting backgrounds. } } } } The variables I am unable to supply are Working Distance, Fixed Distance, } } and Azimuth. I have seen reference to a Quantex Parameters List. This } } document was shipped with the original equipment, but alas, this is a } second } } hand scope and the detector was taken from the company warehouse. } } } } Does anybody use this combination of TEM and detector or know of someone } } with this combination? Does anyone wish to share a document listing } Quantex } } parameters for different scopes with Kevex detectors? } } } } My sanity is in your hands. } } I remain humbly yours, } } } } } } Stephen Bennett } } EMSL Analytical, Inc. } } Miami, FL } } } } miamilab-at-emsl.com } }
I am sure you will find differential staining methods for xylem in there ..
Using Toluidin Blue O gives good results just by the more intense (dark blue) xylem cell walls compared to the thinner (light blue) parenchyma cell walls - but this is not a differential staining method!
Hope this helps you,
Joachim
Dr. Joachim Prutsch Product Manager EM Specimen Preparation
I have a Hitachi S-800 FESEM gun bellows that has developed a small leak. Although it has been replaced, I have heard of bellows being repaired by plating with cadmium or some other metal.
Karli Greg is right. A nuclepore filter could be used to separate small specimens from reagents, which could be exchanged using a syringe. But this is only one of various options. Another option would be to encapsulate the specimens in low-melting point agarose. Then they can be handled in the same way as as tissue blocks. However, centrifugation using a low-speed centrifuge such as an eppendorff centrifuge, will easily separate the unbound gold probe from your specimens. If you are worried about this, test a sample of you colloidal gold probe in your centrifuge. If the supernatant stays pink and there is no red pellet the gold is still a sol. I would recommend that you process the tissue as you are doing, using centrifugation, until the immunolabelling procedure is completed, and at that stage embed the specimen pellet into agarose prior to dehydration and resin infiltration.
Moving your specimen direct from propylene oxide to pure resin is a recipe for specimen collapse unless your specimens are a) exceedingly small, b) very permeable to the resin. What happens is the very mobile PPO comes out of the specimen faster than the viscous resin can move into it, resulting in a volume reduction and shrinkage. You can normally get away with two or three intermediate steps and these are much easier to accomplish if your specimens are in large pieces (we're talking relative sizes here - large means visible, cubes maybe 0.2 to 1 mm in a side) Good luck Chris
----- Original Message ----- } From: "Greg Erdos" {gwe-at-biotech.ufl.edu} To: "Karli Fitzelle" {fitzelle.1-at-osu.edu} ; {Microscopy-at-sparc5.microscopy.com} Sent: Thursday, April 19, 2001 9:58 PM
Hi John,
This is a simple parallax problem.
Imagine the specimen in cross section. If there are two particles one vertically above the other they are seperated by the film thickness T. Tilt the film through an angle A and in plan view the particles will seperate by a distance D. (This can also be extended to account for two particles not vertically above each other but I'll stick to the easy case for the explaination.)
Take two negatives one at zero tilt and one at tilt of A and measure the seperation D, the thickness can be calculated by T=D/sinA.
You will need to know the direction of the tilt axis for your measurements and you can see that the larger the tilt angle and the more acurately you can measure the seperation the more accurate your measurement will be. Tilt + and - A to get a more accurate result.
Now you have to think about what you are going to use to mark the surfaces as you need to be able to distinguish the markers at two different tilt angles. Crystals may go in and out of contrast making them difficult to follow. If you use spheres don't forget to subtract the diameter of the sphere (or one the sum of the two radii) from your total thickness to account for the fact that you are probably using the circumferance as the marker.
Of course the alternative is to re-embed and cross section to measure the thickness directly.
With any luck there may be some responses from people currently measuring thicknesses using this, or other techniques, with some relevant hints or comments on expected accuracy of the different methods.
Good luck, Ron
On Thu, 19 Apr 2001 13:35:43 -0600 John Chandler {chandler-at-lamar.ColoState.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 have a user in our EM Center who would like to have a procedure for } determining the thickness of their ultrathin sections, 60-100 nm. They are } using the normal color criterion of "silver" sections in the boat to select } their sections, but would like to be more precise. There is a potential for } doing morphometry and comparisons of particle counts between non-serial } sections and sections from different specimens. } } They have heard of a technique that uses small particles applied to both } surfaces of the section and using tilt and geometry of the TEM stage to } determine section thickness. Any details of this technique would be } appreciated. } } All suggestions are welcome. } } John } Colorado State University } john.chandler-at-colostate.edu } }
---------------------- 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
We get the bellows replaced by a local (UK) firm, typically around 250 pounds for edge welded bellows in stainless steel. This includes removing the old bellows, supply and welding in new bellows and leak testing. It usually takes a few weeks unless we are prepared to pay to interupt the work scedule. These prices are for 20-40mm dia 50-80mm long bellows.
There must be firms in the US (and most other places) to do this.
} ------------------------------------------------------------------------ } 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 have a Hitachi S-800 FESEM gun bellows that has developed a small leak. } Although it has been replaced, I have heard of bellows being repaired by } plating with cadmium or some other metal. } } Does anyone have any experience with this? } } Thank You, } } Earl Weltmer } } }
---------------------- 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
----- Original Message ----- } From: "Chris Jeffree" {c.jeffree-at-ed.ac.uk} To: {karenco-at-discoverymail.com} Sent: Friday, April 20, 2001 8:52 AM
Howdy Y'all,
I am looking for an inexpensive vise-style specimen holder for our SEM. It does not need to be fancy, just a flat-jawed simple device with a screw to tighten the faces of the jaws. After becoming frustrated by all the information available on the web,I'm sure someone out there could be of assistance. Please help by responding online.
Mr. Weltmer, Try finding an aircraft grade, two-part epoxy (Epoxo 88). If you have stress cracks on a vacuum bellows, mix a batch of epoxy and thinly spread over the joints and areas where you suspect a crack. Make sure to prep the area with acetone or methanol and wait for the solvent to flash off before applying the epoxy. I have repaired a vacuum manifold on a Poloron plasma asher and have not had a leak for a year now. The epoxy doesn't seem to outgass enough to bother. You may want to place the joint in a muffle furnace at low temp. after a few hours to make sure the product is fully cured.
Hi Karen, Here is a paper that deals with immunoelectron microscopy for the SEM. If you have any questions you can contact me at the e-mail or phone below.
Coller, Barry S., Kutok, J. L., Scudder, L. E., Galanakis, D. K., West, S. M . , Rudomen, G. S., Springer, K. T., "Studies of Activated GPIIb/IIIa Receptors on the Luminal Surface of Adherent Platelets: Paradoxical Loss of Luminal Receptors When Platelets Adhere to High Density Fibrinogen." J. Clin. Invest. Vol. 92, pp. 2796-2806, 1993
karenco-at-discoverymail.com wrote: }
} } Email: karenco-at-discoverymail.com } Name: karen chamusco } } Organization: university of florida } } Education: Graduate College } } Location: gainesville, fl } } Question: I am learning to do SEM now and have been doing } immunolocalization (histogold) for light microscopy. I am now } interested in applying this histogold immunolocalization in EM work. } It appears that most immunolocalization I've seen is in TEM. I am } working in SEM. Is there any advantage of one over the other besides } the obvious difference that TEM looks inside the tissue and SEM looks } at the surface? How come there aren't as many SEM localizations out } there (that I've seen)? Thank you for your time, Karen } } ---------------------------------------------------------------------------
-- Regards, Gregory Rudomen Technical Specialist Electron Microscopy State University of New York at Stony Brook University Microscopy Imaging Center Stony Brook, NY 11794-8088 W-631-444-7372 Greg-at-umic.sunysb.edu--http://www.umic.sunysb.edu ************************************************* Standard disclaimer: The opinions expressed in this communication are my own and do not necessarily reflect those of the University Microscopy Imaging Center. *************************************************
The consensus clearly is Duniway Stockroom Corp. of Mountain View, CA. I called them and they have the proper cable in stock for less than half of what Varian wants. Your collective help is greatly appreciated.
If you first remove the plastic from your semi-thin sections, you can use most histological stains. A saturated solution of sodium hydroxide in methanol (sodium methoxide) will remove the plastic from 500 nm sections of plant material in about 10 minutes. Rinse the slides in 100% methanol followed by water and they are ready to go.
Tiny sections are sometimes very hard to find after the resin is removed, so we usually draw around the sections that have been heat-fixed to the slide with a glass scribe.
We (and others) have successfully used analine blue, Auramine O, and Sudan black on Arabidopsis. It is sometimes necessary to remove osmium from the de-plasticized sections (with sodium meta-periodate) for the staining protocols to work.
Most of these techniques are in M.A. Hayat's Principals and Techniques of Electron Microscopy Biological Applications.
If you'd like some references, I'll hunt them up.
Heather Owen
Heather A. Owen, Director Electron Microscope Laboratory Department of Biological Sciences University of Wisconsin - Milwaukee (414)229-6816
Hi Ron, They are robbing you. For less than that price you should get a complete bellow including fittings and freight. That way you can continue operation with the old bellows silicone sealed until the replacment bellows are available. Disclaimer: ProSciTech supplies SS bellows. 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 ABN: 99 724 136 560 www.proscitech.com
On Friday, April 20, 2001 5:53 PM, Ron Doole [SMTP:ron.doole-at-materials.oxford.ac.uk] 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 Earl, } } We get the bellows replaced by a local (UK) firm, typically } around 250 pounds for edge welded bellows in stainless } steel. } This includes removing the old bellows, supply and welding } in new bellows and leak testing. It usually takes a few } weeks unless we are prepared to pay to interupt the work } scedule. } These prices are for 20-40mm dia 50-80mm long bellows. } } There must be firms in the US (and most other places) to do } this. } } Ron } } } On Thu, 19 Apr 2001 22:31:34 -0700 Earl Weltmer } {eweltmer-at-home.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 All, } } } } I have a Hitachi S-800 FESEM gun bellows that has developed a small leak. } } Although it has been replaced, I have heard of bellows being repaired by } } plating with cadmium or some other metal. } } } } Does anyone have any experience with this? } } } } Thank You, } } } } Earl Weltmer } } } } } } } } ---------------------- } 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 }
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Karli Fitzelle writes: I have a quick question. I am working with purified nuclear matrix samples and need tips for the fixation / embedding process.......etc
Hi Karli, Chris Jeffree gave you the advice you need - embed the samples in low melting point agarose. Do this at the very beginning of the process after you have fixed them but washed away the fixative. You can then cut the embedded pieces into small blocks and do all your immunolabeling on these blocks. The agarose is permeable to antibodies and colloidal gold. You may need to increase your incubation and washing times to reflect the reduced accessibility. You will also not have to worry about centrifugation or rushing through the resin embedding as the sample will essentially behave as small tissue pieces.
If you do centrifuge the samples then do not worry about losing the gold - it will remain attached to the antibodies. Centrifuging the gold prior to labeling is another matter.
Regards,
Paul Webster
Paul Webster, Ph.D. Scientist II & Director Ahmanson Advanced Electron Microscopy & Imaging Center House Ear Institute 2100 West Third St. Los Angeles, CA 90057
by ultra5.microscopy.com (8.9.3+Sun/8.9.1) id KAA02466 for dist-Microscopy; Fri, 20 Apr 2001 10:21:44 -0500 (CDT) Received: from no_more_spam.com (sparc5 [206.69.208.10]) by ultra5.microscopy.com (8.9.3+Sun/8.9.1) with SMTP id KAA02463 for "MicroscopyFilteredEmail1-at-msa.microscopy.com"; Fri, 20 Apr 2001 10:21:13 -0500 (CDT) Received: from hera.itg.uiuc.edu (hera.itg.uiuc.edu [130.126.126.145]) by ultra5.microscopy.com (8.9.3+Sun/8.9.1) with ESMTP id KAA02456 for {Microscopy-at-sparc5.microscopy.com} ; Fri, 20 Apr 2001 10:21:02 -0500 (CDT) Received: from buba.uiuc.edu (buba.itg.uiuc.edu [130.126.126.228]) by hera.itg.uiuc.edu (AIX4.3/8.9.3/8.9.3) with ESMTP id KAA257280 for {Microscopy-at-MSA.Microscopy.Com} ; Fri, 20 Apr 2001 10:17:51 -0500 Message-Id: {4.3.1.2.20010420085114.00bd5b40-at-mail.itg.uiuc.edu} X-Sender: gfried-at-mail.itg.uiuc.edu X-Mailer: QUALCOMM Windows Eudora Version 4.3.1
The Imaging Technology Group at the Beckman Institute, University of Illinois at Urbana-Champaign, is seeking a light microscopist to work in our multi-user Microscopy Suite.
Responsibilities will include:
· Operating/maintaining the confocal microscope, fluorescence microscope, stereology workstation, and stereo dissecting microscope. · Supervising and training others in the use of these instruments. · Working in conjunction with users to apply light microscopy techniques to their research. · Developing novel applications to take advantage of the unique capabilities of this instrumentation.
A complete job description is posted on the ITG web page http://www.itg.uiuc.edu or http://www.itg.uiuc.edu/ms/lightmicroscopistjob.pdf
Please send a letter of application and resume to:
Lori Heil Imaging Technology Group Beckman Institute for Advanced Science and Technology 405 N. Mathews Urbana, IL 61801 (217) 244-0170 e-mail: lheil-at-uiuc.edu
The University of Illinois is an Affirmative Action/Equal Opportunity Employer. Women and minorities are encouraged to apply.
Ray, I have made a number of specimen holders over the last few years, at rock- bottom prices :). My recipe for a pin-mount style mini-vice is: - grind or cut a small block of brass to the vice size desired - drill one hole in the large face of the block the size of the pin (typ. 3mm) - drill two parallel holes (the size for tap holes) through the long axis of the block - cut off one end of the block perpendicular to the two tap holes - tap the holes in the larger section of the block - drill out the tap holes in the smaller section of the block to make clearance holes - cut off a length of brass rod for the pin mount, and solder it in place in the hole in the larger block - screw two bolts (brass recommended) into the large block through the clearance holes in the smaller block. This is your mini-vice. - de-burr, clean, and polish as desired.
With proper tools several of these can be made in an hour; with what's available from the local hobby store, maybe 1.5-2 hours each.
Ben (simkin-at-egr.msu.edu)
} Howdy Y'all, } } I am looking for an inexpensive vise-style specimen holder for our } SEM. It does not need to be fancy, just a flat-jawed simple device } with a screw to tighten the faces of the jaws. After becoming } frustrated by all the information available on the web,I'm sure } someone out there could be of assistance. Please help by responding } online. } } Regards, } } Ray Grassl } } Grassl.raymond-at-basco.com
} } } } } } I have a user in our EM Center who would like to have a procedure for } } determining the thickness of their ultrathin sections, 60-100 nm. They are } } using the normal color criterion of "silver" sections in the boat to select } } their sections, but would like to be more precise. There is a potential } } for } } doing morphometry and comparisons of particle counts between non-serial } } sections and sections from different specimens. } } } } They have heard of a technique that uses small particles applied to both } } surfaces of the section and using tilt and geometry of the TEM stage to } } determine section thickness. Any details of this technique would be } } appreciated. } } } } All suggestions are welcome. } } } } John } } Colorado State University } } john.chandler-at-colostate.edu } }
The is a very good review of measuring section thickness in Audrey Glauert's book "Sectioning and Cryosectioning for Electron Microscopy". Several different methods are discussed. In short, it is difficult to know precisely how thick a silver section is. My edition of the book was published in 1991. There may be newer and more encouraging reports that I am not aware of.
Good Luck,
John
John M. Basgen Department of Pediatrics University of Minnesota Mayo Mail Code 491 420 Delaware Street SE Minneapolis, MN 55455 USA Phone: 612-625-7979 FAX: 612-626-2791 E-mail: basgen-at-umn.edu
Karli, have you considered using magnetic beads to carry your organelles through these steps? The process is less disruptive to fragile organelles than centrifugation. Whether the gold will settle during centrifugation will depend on the size of the gold, an example is 15 nm. gold settles at 25,000 rpm for 60 minutes. For a discussion of this issue, you can consult the early papers describing the conjugation of various proteins to colloidal gold, Marge
Margaret Springett IEM Specialist Electron Microscopy Core Facility Mayo Foundation email: springett.margaret-at-mayo.edu
} ---------- } From: Karli Fitzelle } Sent: Thursday, April 19, 2001 2:27 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: TEM Question
In a message dated 4/19/2001 3:01:22 AM Mountain Daylight Time, RCHIOVETTI-at-aol.com-at-sparc5.microscopy.com writes:
} Does anyone know of a digital imaging system for scanning EM?
A list of companies that provide digital imaging systems for SEMs can be found at "www.jcnabity.com/links.htm#Digital Imaging".
The two basic types of systems are passive (where they acquire an image while the SEM scans) and active (where they control the beam position during the image acquisition). The passive types will typically be less expensive, since they do not need to generate the sweep voltages. The active systems require that the SEM has XY inputs for beam control.
Joe _________________________________________ Joe Nabity, Ph.D. JC Nabity Lithography Systems E-Beam Lithography using Commercial SEMs & STEMs PO Box 5354, Bozeman, MT 59717 USA Voice: (406) 587-0848 FAX: (406) 586-9514 E-mail: info-at-jcnabity.com Web: www.jcnabity.com
I teach an undergraduate-level course in electron microscopy, and every year I find that the biggest hurdle for my students, not surprisingly, is the cutting of thin sections with glass knives. This is of course the point at which the students begin to get very discouraged with their individual research projects. I have tried various embedding media based on viscosity/penetration demands (Spurr's or ultra-low viscosity for plant tissue, for example), but I have been unable to come up with a medium that gives students a greater chance at success in cutting sections with glass knives. Does anyone have a favorite embedding medium that would allow fairly routine sectioning of diverse biological samples with glass knives. We are using primarily MT-2 microtomes.
Earl Weltmer wrote: ============================================= I have a Hitachi S-800 FESEM gun bellows that has developed a small leak. Although it has been replaced, I have heard of bellows being repaired by plating with cadmium or some other metal. ============================================== Is this one of those kinds of leaks that could be repaired, at least temporarily, with a product like VacSeal™ vacuum leak sealant? See URL http://www.2spi.com/catalog/vac/vacleak.html
It is used even on UHV systems for "temporary" repairs, but depending on the nature of the leak, sometimes these "temporary" repairs can last months or even years.
And this kind of fix is quick and cheap!
Disclaimer: SPI Supplies offers this product to those with vacuum leaks.
Chuck
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Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
I'd like to prepare some TEM films of 30-50 Angstrom CdS particles that are in a block-co-polymer solution. Has anyone done this? Could we just pour them onto a carbon film and let it dry to examine the particles? Any other ideas?
I think that glass knives dull very rapidly (either damage or plastic build-up on the edge), and so we have always tried to get the first few sections at a given place on the knife edge, which tend to be the best. I wrote a brief procedure for students years ago, based on that approach, and have typed it below (the handout had a drawing of a glass knife, with area A = 1/3 of edge on the side where the whorl meets the edge, area B = middle 1/3 of knife edge, area C = 1/3 of edge at side where whorl is farthest from the edge):
1. Face the block in area C of knife (see diagram). Cut semithin sections (if desired) in area B of knife. Then move to area A.
2. Bring the block face parallel with the knife, using the shadow method. Then use the shadow to bring the block face as close to the knife (but without touching) as you dare.
3. With the ultramicrotome set for ultrathin sections, manually turn the microtome wheel quite rapidly until you see the first sign of contact (usually a sliver off one side of the block face), then stop turning.
4. Turn on automatic sectioning at usual slow cutting speed. Cut about 6-12 full-face sections, then stop and pick them up on EM grids.
5. Retract the stage slightly, move laterally to another place in area A, and repeat steps 2-4. Continue until area A has all been utilized (or until you have all the sections you need).
Good luck to you and your students.
Kent
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ A. Kent Christensen, Professor Emeritus Department of Cell and Developmental Biology, Medical Science II Building University of Michigan Medical School, Ann Arbor, MI 48109-0616 Office: 5801 Medical Science II Building Tel (work) (734) 763-1287, Fax (work) (734) 763-1166 E-mail: akc-at-umich.edu ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
} I teach an undergraduate-level course in electron microscopy, and } every year I find that the biggest hurdle for my students, not } surprisingly, is the cutting of thin sections with glass knives. } This is of course the point at which the students begin to get very } discouraged with their individual research projects. I have tried } various embedding media based on viscosity/penetration demands } (Spurr's or ultra-low viscosity for plant tissue, for example), but I } have been unable to come up with a medium that gives students a } greater chance at success in cutting sections with glass knives. } Does anyone have a favorite embedding medium that would allow fairly } routine sectioning of diverse biological samples with glass knives. } We are using primarily MT-2 microtomes. } } I thank you in advance. } } Dick Briggs } Biology Department } Smith College
Karli, have you considered using magnetic beads to carry your organelles } through these steps? The process is less disruptive to fragile organelles } than centrifugation. Whether the gold will settle during centrifugation } will depend on the size of the gold, an example is 15 nm. gold settles at } 25,000 rpm for 60 minutes. For a discussion of this issue, you can consult } the early papers describing the conjugation of various proteins to colloidal } gold, } Marge
} Margaret Springett } IEM Specialist } Electron Microscopy Core Facility } Mayo Foundation } email: springett.margaret-at-mayo.edu
} } ---------- } } From: Karli Fitzelle } } Sent: Thursday, April 19, 2001 2:27 PM } } To: Microscopy-at-sparc5.microscopy.com } } Subject: TEM Question
Robin, Why don't you try polymerizing the block co-polymer and then microtoming the sample. You could get someone over in the med school to do it for you. There are two places on campus that have the facilities and expertise to do it for you, biology, and pathology. The size of the particles would make it very straightforward to do. Talk to Jim Sheetz.
-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: "rgriffin-at-eng.uab.edu"-at-sparc5.microscopy.com } [mailto:"rgriffin-at-eng.uab.edu"-at-sparc5.microscopy.com] } Sent: Friday, April 20, 2001 2:42 PM } To: microscopy-at-sparc5.microscopy.com } Subject: Sample prep of CdS particles in block co-polymer mycells } } } -------------------------------------------------------------- } ---------- } 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/MicroscopyLists } erver/FAQ.html } } } } -------------------------------------------------------------- } ---------. } } } I'd like to prepare some TEM films of 30-50 Angstrom CdS } particles that are } in a block-co-polymer solution. Has anyone done this? Could } we just pour } them onto a carbon film and let it dry to examine the } particles? Any other } ideas? } } } Thanks, } } } Robin Griffin } UAB }
I worked in an MBE group where we grew epitaxial and heteroepitaxial layers on III-V compounds. I thought that I had a pretty good idea of what epitaxy is.
Recently, I have come across a rash of papers that claim epitaxial relationships across dissimilar materials with dissimilar crystal structures. This is just orientational relationships across the interface. It is not epitaxy! Is it because we as materials scientists/microscopists who know better are not getting to review these papers or has there been a change of the definition of epitaxy. What's going on? (This is a rhetorical question posed to foster a discussion.) What are we going to do about it? (another one.) Am I the only one that is concerned that the definition is becoming unclear?
I know that I have made comments about it when I review papers that claim epitaxy when it is really just an orientational relationship.
-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
I am looking for a small oven in which to polymerize epoxy embedded samples (50-90 C). I'd like one small enough to fit into our fume cabinet, but the smallest one I can find is about 13 x 13 x15 inches. Does anyone know of a source for a smaller oven?
Thanks in advance,
Doug ---------------------- Douglas R. Keene Associate Investigator Shriners Hospital Research Facilities 3101 S.W. Sam Jackson Park Road Portland, Oregon 97201 phone: 503-221-3434 FAX: 503-412-6894 (9-5 PST) e-mail: DRK-at-shcc.org
Does LKB still manufacture glass strips? If so, from whom does one obtain such glass? If not, what is the best glass available for sectioning and an approximate price? I vaguely remember "LKB glass" and have been using an unspecified "brand X". Thank you for your replies.
Barbara Plowman Univ. of the Pacific School of Dentistry 2155 Webster St. San Francisco, CA 94115 email: Bplowman-at-sf.uop.edu ph: 415-929-6692
You might want to try contacting Stuart Enterprises run by Stuart Wisun. Specimen holders for SEMs are his specialty. His number is (650)424-9089, and his email address is stuwsn-at-juno.com.
Tom Pella
Raymond Grassl 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 } -----------------------------------------------------------------------. } } Howdy Y'all, } } I am looking for an inexpensive vise-style specimen holder for our } SEM. It does not need to be fancy, just a flat-jawed simple device } with a screw to tighten the faces of the jaws. After becoming } frustrated by all the information available on the web,I'm sure } someone out there could be of assistance. Please help by responding } online. } } Regards, } } Ray Grassl } } Grassl.raymond-at-basco.com
LKB is now part of Leica, but LKB never was a glass manufacturer. They marketed that glass only. Its been sold under the manufacturer's brand name "Alkar" for many years now. All EM suppliers sell that glass. We do sell Alkar glass in Australia and New Zealand, but clearly its one item that you would buy somewhere on your continent. 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 ABN: 99 724 136 560 www.proscitech.com
On Saturday, April 21, 2001 8:40 AM, Barbara Plowman [SMTP:Bplowman-at-sfmail.dental.uop.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 } -----------------------------------------------------------------------. } } } Does LKB still manufacture glass strips? If so, from whom does one obtain } such glass? If not, what is the best glass available for sectioning and an } approximate price? I vaguely remember "LKB glass" and have been using an } unspecified "brand X". Thank you for your replies. } } Barbara Plowman } Univ. of the Pacific } School of Dentistry } 2155 Webster St. } San Francisco, CA 94115 } email: Bplowman-at-sf.uop.edu } ph: 415-929-6692 }
Robin Griffin wrote: ============================================================= I'd like to prepare some TEM films of 30-50 Angstrom CdS particles that are in a block-co-polymer solution. Has anyone done this? Could we just pour them onto a carbon film and let it dry to examine the particles? Any other ideas? ============================================================== What tends to happen is that the polymer is present in sufficient amount that it interferes with the imaging. Hence you can get around this problem by putting your preparation on silicon monoxide/dioxide filmed grids, then in an oxygen plasma etcher, etch away the polymer, leaving only the CdS colloid dispersed on the support film. However it might be better to put your solution on a glass slide, allow it to dry, and then etch off the organics as above, then Pt/C shadow or carbon coat only and pick up the replica on a grid and examine.
Disclaimer: SPI Supplies manufactures a plasma etcher and also produces SiO2 filmed grids.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
Epitaxial growth is the oriented growth of a crystalline material over another crystalline material [9]. For example, films of fcc Au{001} can be grown by the deposition of gold on fcc Ag{001} surfaces; the silver lattice-parameter (4.08 Ang.) is very close to the gold lattice-parameter (4.09 Ang.). In some case the meshes may not match, e.g., Tb(0001) grown on W{110}, but incommensurate growth is still possible. In other cases, the meshes may match by the expansion, dilation, and rotation of the meshes, as Pb{111} grown on Si{111}. The success or failure of epitaxial growth depends highly upon the chosen material's chemical and physical properties, as well as the surface structure of the substrate.
} From: "Walck, Scott D." {walck-at-ppg.com} } To: "'Microscopy'" {microscopy-at-sparc5.microscopy.com} } Subject: Epitaxy -Am I misinformed or what? } Date: Fri, 20 Apr 2001 17:59:41 -0400 } } ------------------------------------------------------------------------ } 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 worked in an MBE group where we grew epitaxial and heteroepitaxial layers on III-V compounds. I thought that I had a pretty good idea of what epitaxy is. } } Recently, I have come across a rash of papers that claim epitaxial relationships across dissimilar materials with dissimilar crystal structures. This is just orientational relationships across the interface. It is not epitaxy! Is it because we as materials scientists/microscopists who know better are not getting to review these papers or has there been a change of the definition of epitaxy. What's going on? (This is a rhetorical question posed to foster a discussion.) What are we going to do about it? (another one.) Am I the only one that is concerned that the definition is becoming unclear? } } I know that I have made comments about it when I review papers that claim epitaxy when it is really just an orientational relationship. } } } -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) } } }
Since Joe's list for some reason unknown to me does not include my company, I would also like to invite you to take a look at our web site. We provide both passive and active systems.
Thanks.
Michael Bode Soft Imaging System Corp. www.soft-imaging.com mb-at-soft-imaging.com
-----Original Message----- } From: "NPGSlithography-at-aol.com"-at-sparc5.microscopy.com [mailto:"NPGSlithography-at-aol.com"-at-sparc5.microscopy.com] Sent: Friday, April 20, 2001 10:13 AM To: RCHIOVETTI-at-aol.com; Microscopy-at-sparc5.microscopy.com
In a message dated 4/19/2001 3:01:22 AM Mountain Daylight Time, RCHIOVETTI-at-aol.com-at-sparc5.microscopy.com writes:
} Does anyone know of a digital imaging system for scanning EM?
A list of companies that provide digital imaging systems for SEMs can be found at "www.jcnabity.com/links.htm#Digital Imaging".
The two basic types of systems are passive (where they acquire an image while the SEM scans) and active (where they control the beam position during the image acquisition). The passive types will typically be less expensive, since they do not need to generate the sweep voltages. The active systems require that the SEM has XY inputs for beam control.
Joe _________________________________________ Joe Nabity, Ph.D. JC Nabity Lithography Systems E-Beam Lithography using Commercial SEMs & STEMs PO Box 5354, Bozeman, MT 59717 USA Voice: (406) 587-0848 FAX: (406) 586-9514 E-mail: info-at-jcnabity.com Web: www.jcnabity.com
Hi Dick Briggs: I was a former student at San Francisco State University (I've graduated in May 2000), and I took an electron microscopy course at that university. My instructor was Dr. Greg Antipa, and I'm sure that he can give you tips on plastics he used, etc. Back to my electron microscopy course ... during part of that course, I was supposed to section some embedded tissue of your choice (heart / liver / ? (can't recall off-hand)), and I had considerable trouble using a microtome well. As I recall, we had some Sorvalls available for use, but I'm unsure if they were MT-2s. I prepared all my tissues (and practice, plastic "pellets") by first cutting with straight razor blades to make an acceptable-sized pyramid, then continue by sectioning with a microtome. Glass knives were used on these embedded tissues -- the plastic used was Epon -- and I recall that one major problem I had was in making sure that the microtome sectioned at a consistent speed. In addition, the glass knife edges would dull pretty quickly after a small number of sections were made, and so it became imperative to use several glass knives for any particular embedded material. I eventually was able to section pretty well, but I also picked up sections from below with a grid (of 200 mesh) that I bent rather badly during the collection process. Epon worked OK for sectioning, and while it's true that students can indeed get discouraged, I also know from experience that students have very different aptitudes towards handling microtomes, preparing the pyramids, etc. I hope that my experience with sectioning may help you understand better (from a student's perspective) the various trouble areas that could occur when students are first learning how to section embedded material. Good luck with the sectioning process. Nelson Conti
_______________________________________________________ Send a cool gift with your E-Card http://www.bluemountain.com/giftcenter/
On Monday April 23rd 2001 at ~ 9AM CST , after more than two decades of operation, the ANL HVEM will begin its last experimental session before permanently shutting down at the end of the day.
On behalf of the hundreds of users in the world wide microscopy research community who have used this facility since 1979, let me offer thanks to the ANL crew who has kept this unique resource alive, functional and running over this period: Hats off to ...Ed Ryan, Stan Ockers, Charlie Allen, Tony McCormick, as well as Alan Philippides, Loren Funk, Loren Thompson, Pete Baldo, and Tony Taylor.
If you would like to peer "live" into the microscope room via TelePresence and be a small part of the last day of operation of this unique instrument for Materials Research just point your Web browser to
http://tpm.amc.anl.gov/HVEM.html
When the last experiment completes later the afternoon of April 23rd, ANL will begin the decommissioning process and over the next few weeks the instrument will be completely dismantled. During this time we will endeavor to keep the live telepresence links operational to broadcast the decommissioning of this unique resource to allow interested students/researchers the opportunity to observe the process.
Coincidentally the last experiment being conducted on the HVEM will be a high energy electron irradiation damage study similiar in many respects to the first official experiment conducted at the facility during its opening ceremonies over twenty years ago.
Nestor J. Zaluzec Materials Science Division Argonne National Laboratory
-- =========================================== Dr. Nestor J. Zaluzec Materials Science Division Building 212 Argonne National Lab 9700 S. Cass Ave Argonne, Illinois 60439 USA Tel: 630-252-7901, Fax: 630-252-4289 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 !
Does anyone know how to detect glycolipid using LR-White/Cryo ultrathin-sections? I have tried both. But in cryosections the lipid is flowing out over the section, so when I do my immunogold, there is gold all over. I think that I lose some lipid bound to the membrane, when I use LR-White. Maybe because of the dehydrasion. Maybe there is a speciel way to fix the lipid?
The Biomaterials Science Group Department of Oral and Dental Science University of Bristol in collaboration with Glaxo SmithKline
Postdoctoral Research Assistant
in the Biomaterials Science Group on the project Interaction Mechanisms of Polymers at Interfaces of Mineralised Tissues
The research area involves the study of the physical and chemical properties and interaction mechanisms of different polymers at interfaces of mineralised tissues. You will have recently been awarded a PhD in an appropriate field and will ideally have experience in scanning probe microscopy (AFM) of biological materials and other analytical techniques and an interest in medical research. You will work in Dr. Jandts group and interact with scientists at Glaxo Smith Kline. The University of Bristol is one of the leading research universities in the UK and provides an outstanding scientific training environment to enhance your qualification. The group is involved in exciting, interdisciplinary projects and maintains appropriate state of the art instrumentation. There exist opportunities for additional interactions with clinical scientists and other centres at the university. We are looking for a dynamic and exceptionally well-qualified postdoctoral researcher who can interact effectively in an international and interdisciplinary team. The appointment will be on a Research Assistant 1A scale with a salary range of # 16775 to # 20465. This is a full time appointment and initially for one year. Applicants should include a short CV, stating research experience and interests, publication list and addresses of two referees. The review of applications will start 24 May 2001 and will continue until the post has been filled. Informal inquiries can be directed by email to Dr. K. D. Jandt (K.Jandt-at-bris.ac.uk), Senior Lecturer in Biomaterials, University of Bristol
Formal applications quoting the reference number 7401 should be directed to
The University of Bristol Recruitment Office Bristol, BS8 1TH United Kingdom
----------------------------------------------------------------- Dr. rer. nat. Klaus D. Jandt Senior Lecturer in Dental Materials Science and Biomaterials University of Bristol, Department of Oral and Dental Science Lower Maudlin Street, Bristol, BS1 2LY, UK Phone: +44-117-9284418, Fax: ++44-117-9284780 Internet: K.Jandt-at-bris.ac.uk WWW: http://www.dent.bris.ac.uk/Biomaterials/kdj.htm "We make Biomaterials Science work!"
Does anyone know how to detect glycolipid using LR-White/Cryo } ultrathin-sections? } I have tried both. But in cryosections the lipid is flowing out over the } section, so when I do my immunogold, there is gold all over. } I think that I lose some lipid bound to the membrane, when I use LR-White. } Maybe because of the dehydrasion. } Maybe there is a speciel way to fix the lipid?
What a sad morning this is. I've used this instrument, way back around 1980, and I can say it's a crying shame that such machines have such short lives. I found it to be quite useful for my area of study (finding the orientation distribution, more properly the misorientation distribution, in the cellular substructures of highly cold worked metals) for the simple reason that the selected area aperture of a high-voltage microscope can be demagnified sufficiently accurately for the electron beam to be made to illuminate the sub-micron-sized crystallites.
I've outlived _two_ high-voltage microscopes. Before the ANL instruments, I was a heavy user of the Million-Volt TEM at the US Steel Research Center, and for the same reason. Unfortunately, I was driven to use Argonne's HVEM because US Steel's hourly charges were too high to fit in my research budget; what with lack of support for what should have become a regional resource, the US Steel instrument was soon decommissioned just like ANL's is about to be.
Best regards, George Langford, Sc.D. amenex-at-amenex.com http://www.amenex.com/
Hi Scott, I think the term 'epitaxy' has indistinct boundaries. Okay, so InGaAs on GaAs is undisputedly epitaxy, as long as the crystals line up. But what about GaAs on Si? Most people would say this is epitaxy, but they're different crystal structures (diamond on sphalerite). Would you say something with rock salt structure can't be epitaxial with a diamond structure substrate? And just to stretch it a bit further, how about a (111) cubic layer on a (0001) hexagonal substrate? If not, what about hexagonal CdS on cubic CdS? My interpretation is that epitaxy requires both a reasonably fixed orientation relationship and a deposition. So MBE silicon on sapphire can be epitaxial, but a martensitic phase transformation can not. Maybe I should look it up in a science dictionary...
Richard
} ------------------------------------------------------------------------ } 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 worked in an MBE group where we grew epitaxial and heteroepitaxial layers on III-V compounds. I thought that I had a pretty good idea of what epitaxy is. } } Recently, I have come across a rash of papers that claim epitaxial relationships across dissimilar materials with dissimilar crystal structures. This is just orientational relationships across the interface. It is not epitaxy! Is it because we as materials scientists/microscopists who know better are not getting to review these papers or has there been a change of the definition of epitaxy. What's going on? (This is a rhetorical question posed to foster a discussion.) What are we going to do about it? (another one.) Am I the only one that is concerned that the definition is becoming unclear? } } I know that I have made comments about it when I review papers that claim epitaxy when it is really just an orientational relationship. } } } -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) } }
e-mail richard.beanland-at-marconi.com Tel. +44 1327 356363 Fax. +44 1327 356398 ============================================================== "The information contained in this message is legally privileged and confidential information intended for the eyes of the individual or entity named above. If the reader of this message is not the intended recipient, you are hereby notified that any dissemination, distribution or copying of this message is strictly prohibited. If you have received this message in error, please notify us immediately by telephone. Caswell Technology is the trading name of Marconi Caswell Limited. Registered in London No. 3694360 Registered Office: One Bruton Street London W1X 8AQ. Holding Company: Marconi plc."
We are planning to do some 3D reconstruction of tick salivary glands from 2D light microscope sections (1 micron thick).
Does anyone have any suggestions for software (PC) that could do this.
thanks
Kevin
Kevin Mackenzie Electron Microscope unit Department Zoology University of Aberdeen Tillydrone Avenue Aberdeen AB24 2TZ ----------------- Tel 01224 272847 Fax 01224 272396 email k.s.mackenzie-at-abdn.ac.uk Web Site http://www.abdn.ac.uk/emunit
In June, I will have a mid-level EM Tech position open. One of my folks is getting married and moving away. We do all TEM--a mixture of negative staining and thin sectioning; 75% is clinical (diagnosing viral diseases with the EM, plus surgical pathology EM) and 25% is research. We also do some teaching of medical and graduate students and residents. We have 5 tech positions, and the 4 remaining individuals are all delightful folks with whom to work. Durham, NC, is a pleasant place in which to live and work--3 hrs from the mountains or the beach, and has more entertainment than you could want with 3 major universities and their arts programs in close proximity--not to mention the NCAA champs (Go Duke!).
Duke requirements for the position (EM Technician, Senior) include a bachelorąs degree and electron microscopy training (a course or experience) or an associateąs degree in EM. I am looking for someone who is proficient in cutting thin sections and has experience running an electron microscope. We can teach you to operate the particular scope brand we have, including digital operation for some applications, and we can teach negative staining and virus recognition. I am also looking for someone who enjoys challenging and interesting cases, is dedicated to accuracy, and is willing occasionally to put in extra effort in return for appropriate compensation and consideration when you have special needs. And I am particularly looking for someone who can manage several jobs at once while having a good time sharing camaraderie with the rest of us, i.e., is not high strung. I know this special person exists, since there are 4 lovely folks remaining (3 guys and 1 gal) with these same qualifications. I will be happy to answer any questions you have by phone or email.
If you might be interested, please contact me directly as the position is not officially open yet; I have requested that the paperwork be started. My address and phone number follow.
Sara E. Miller, Ph. D. P. O. Box 3712 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-3265
Hi, I need advice on embedding Thermanox coverslips. It's supposed to peel off leaving the monolayer behind, but I'm not having much luck. I'm using Aralidite 502 as the embedding medium. Would a another resin work better? I would appreciate any and all advice. Thanks.
Tom Bargar EM Lab UNMC 402-559-7347 tbargar-at-unmc.edu
A thread on plant cell wall staining for LM reminded me of a project wherein I would have liked to differentially stained parts of the plant cell wall - but for TEM. I could not readily find a good protocol for a CHO EM "stain" to do this. Does anyone have some good protocols and ideas for adding contrast or staining the plant cell wall?
I would like to express my heart felt sadness that another functioning HVEM will be decommissioned today. As one who has spent over 20 years working with almost the same microscope here in Albany, NY it is especially difficult to watch this days events. I know these microscopes like many know the inner workings of good vintage cars or the DC-3 and its difficult for me knowing how successfully the the ANL crew has been throughout the years with this very servicable machine.
There are 6 and then there 5 HVEMs in the US five years ago. Now there will be only 4!
I'm sure just like today we and NASA marvel at how quickly the technology and support for the Saturn 5 technology has disapeared even as our space program is just getting off low earth orbit, we in the microscopy field will find in the very near future how impossible it is to go back to the highvoltage technology that gave us that special edge for thick and dense specimens.
Dave
David Barnard HVEM Wadsworth Center NYS Dept Health Albany,NY
Autoquant's Autovizualize-3D will do this nicely. The resulting 3D image may be rotated and tilted and undergo further processing if desired.
Please contact me off-line if you are interested in this product or need assistance.
Gary Gaugler, Ph.D. Optical Reflections 916.791.8191 916.791.8186 7970 Twin Rocks Rd Granite Bay, CA 95746-8111 USA
Disclaimer: I am an authorized reseller of Autoquant's image processing software products. I mostly handle the Western US. If you have problems obtaining information on Autoquant products in the UK, please contact me for assistance.
See http://www.aqi.com for product technical info.
At 07:09 AM 4/23/2001, you wrote:
} Dear All } } We are planning to do some 3D reconstruction of tick salivary glands } from 2D light microscope sections (1 micron thick). } } Does anyone have any suggestions for software (PC) that could do this. } } thanks } } Kevin } } } Kevin Mackenzie } Electron Microscope unit } Department Zoology } University of Aberdeen } Tillydrone Avenue } Aberdeen } AB24 2TZ } ----------------- } Tel 01224 272847 } Fax 01224 272396 } email k.s.mackenzie-at-abdn.ac.uk } Web Site http://www.abdn.ac.uk/emunit
We still run a Kevex Quantum on a JEOL 840A SEM. We replaced the Delta V portion of our analyzer a few years ago, but still have the old chassis sitting here if you (or anyone else) ever need parts.
I believe most of these concepts were explained in either the Quantex manual (pg. G-65 for version 5)or in the Tutorial (pg.8-8) manual.
Working distance on an SEM was the distance from pole piece to sample surface. Fixed distance was the distance from the pole piece to the centerline of the detector crystal. Therefore, the combination of (WD-FD) and HD was used to calculate takeoff angle. Therefore, if you cannot get the exact numbers for your scope, you can probably measure the height difference between sample and detector and make up some half-reasonable numbers that generate the correct difference.
Azimuth angle fixed the detector position on the column. Assuming the sample holder tilts about the y-axis, azimuth was the angle between x-axis and the detector port. Thus, an azimuth angle of 0 degrees would mean that your sample tilts directly toward the detector. Our detector was located 23 degrees back of the x-axis. If you never tilt your sample, then azimuth would not matter. If you do, it helps to correct solid angle and takeoff angle for the effects of tilt.
We had a tilted detector on our 840. That made things a little trickier. We had a scale reading to show the distance from detector to sample. But since our detector was tilted, we could not use that straightaway for horizontal distance (HD). We had to look up FD and HD values from a table as a function of scale reading. It was a fairly simple exercise in geometry, but it always seemed strange that fixed distance (FD) was not really fixed when the detector was tilted.
I hope this gets you going. If not, just ask for clarification.
Warren
At 09:30 PM 4/19/2001 -0500, you wrote: } } Subject: EDXA, Need help with detector geometry } } } Sirs or Madams, } } } } } } I am running a JEOL CX II with a Kevex mod. 3200-0018 detector/ Kevex } Delta Class Anlyzer. } } } } } } I am having difficulty locating the geometric variables unique to this } } } mating of scope and detector. Kevex was unable to supply the data. } These } } } geometric variables are used by the analyzer software (Quantex) in } modeling and subtracting backgrounds. } } } } } } The variables I am unable to supply are Working Distance, Fixed Distance, } } } and Azimuth. I have seen reference to a Quantex Parameters List. This } } } document was shipped with the original equipment, but alas, this is a } } second hand scope and the detector was taken from the company warehouse. } } } } } } Does anybody use this combination of TEM and detector or know of someone } } } with this combination? Does anyone wish to share a document listing } } Quantex parameters for different scopes with Kevex detectors? } } } } } } My sanity is in your hands. } } } I remain humbly yours, } } } } } } Stephen Bennett } } } EMSL Analytical, Inc. } } } Miami, FL } } } } } } miamilab-at-emsl.com } } ---------------------- } Warren E. Straszheim } Materials Analysis and Research Lab } Iowa State University } 23 Town Engineering } Ames IA, 50011-3232 } } Ph: 515-294-8187 } FAX: 515-294-4563 } } E-Mail: wesaia-at-iastate.edu } Web: www.marl.iastate.edu } } Scanning electron microscopy, x-ray analysis, and image analysis of materials } Computer applications and networking
Job Posting for Electron Bioimaging Lab, submitted by David P. Bazett-Jones
Service Manager, Electron Microscopy Facility
Date Posted: April 17, 2001
Position Status: Full-time, Fixed term
Department: Cell BiologyResearch Institute
Available: August 1, 2001
Description of the Position: You will share responsibility for the operation and maintenance of transmission and scanning electron microscopes in a new Bioimaging Facility co-sponsored by teaching hospitals in the University of Toronto. The microscopes include an ESEM (FEI/Philips) and a 200 kV TEM (FEI/Philips) equipped with EDX, GIF and cryostage. You will also be responsible for coordination and management
of electron bioimaging services required by investigators of the Hospital for Sick Children Research Institute. Qualifications: As an ideal candidate, you have completed a M.Sc. in biological sciences, or have completed a B.Sc. with experience in analytical electron microscopy, ultramicrotomy and other sample preparation techniques. Strong computer skills are an asset.You possess excellent verbal communication andorganizational skills. You have the ability to work well independently and in a team.
Hours : 35 hours/week
Salary: $39,848.95 - $50,277.67
Available to: Internal and External Candidates
Deadline: April 25, 2001
Submit Resume to : Erin O’HareThe Hospital for Sick Children, 555 University Avenue, Toronto, OntarioM5G1X8 Fax (416) 813-5671 E-mail: hr.recruiter-at-sickkids.on.ca
Must Quote File Number CG0102-EO We thank you in advance for your interest. Only those applicants selected for an interview will be contacted.
Hello Tom, I use Thermanox coverslips a lot here, so here is my advice: I always place the coverslip inverted on a drop of resin (if I don't, the coverslip will be "embedded" in the resin and is really hard to remove) on an aclar sheet. I have also found that the small weigh dishes, ours are white, are great for keeping many coverslips separate. Just put one coverslip per dish. They do tend to migrate on the aclar sheet, especially when the oven isn't level. Usually they peel off very easily. In the rare instance that they don't, I use a dissecting scope and choose the area I am interested it. I then use a razor blade and cut through the resin side and remove the area of interest. The small piece will come off nicely and is the exact size and shape I need to remount it on a "blank" block for sectioning. This is also great because the rest of the sample (cell
culture) remains whole and labelled for identification later. I hope this helps, Jo Dee
-- Jo Dee Fish Coordinator of Electron Microscopy Cell Analysis Facility The Burnham Institute 10901 N. Torrey Pines Rd. La Jolla, CA 92037 (858)646-3100 ext. 3620
Does anyone know if NaNO3 and KNO3 are or are not reasonably stable under an electron beam?
I want to use them as overlap standards for Na and K respectively, but would prefer to avoid explosions in the specimen chamber. They would be mounted on conductive double-sided tape, and carbon coated, so there is some oxidisable matter available.
Anybody been there?
cheers
rtch
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
To those biologist out there in cyber space who may have used monastral blue as a marker for macrophages years ago. Many years ago I purchased 3% monastral blue (copper phalocyanine) in solution from Sigma which we used for in vivo and in vitro phagocytic experiments. Unfortunately Sigma now only sell the powder form of monastral blue. Is there any one who has an old bottle of Sigma 3% monastral blue solution that can tell me what was the solvent used to prepare this solution so that I can dissolve this dye to obtain a 3% stock solution. I have tried every solution that I have in the laboratory to dissolve the powder without success. I need to know exactly what the solvent was in the old Sigma solution. Hope someone out there can help.
Terry Robertson (PhD)
Dr Terry Robertson (PhD) Senior Research Fellow Department of Pathology University of Western Australia Nedlands 6009
The best way to clean immersion oil from a lens is to use lighter fluid. First remove as much oil from the lens with lens paper. Be gentle; don't rub. Then dip a cotton swab in the lighter fluid and lightly wipe it across the lens. All the oil will be removed and it will evaporate very quickly without leaving a residue or streaks. Xylene is not recommended as it can dissolve the adhesives holding the lens in place.
Cheryl Rehfeld Meyer Instruments, Inc. Leica Distributor -----Original Message----- } From: mckaylodge-at-aol.com {mckaylodge-at-aol.com} To: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com}
Consider this option to an actual oven: I've used a TEMP-BLOK MODULE HEATER (by Lab-Line, distributed by Scientific Products) for curing resins. Its a small heater measuring about 5x8x3 inches, has high and low temperature ranges with variable control for each. They have a variety of removable blocks with arrays of wells in them into which you can put BEEM capsules, Eppendorf tubes, gelatin capsules,etc. I stick a thermometer in one of the wells to calibrate the tepmerature settings. Place a tight covering of aluminum foil over the top of the block to better stabilize the temperature inside.
I have bought them used from our University's scientific apparatus shop which trades in used lab gear.
Good luck,
Gib
} } Dear Microscopists, } } I am looking for a small oven in which to polymerize epoxy } embedded samples (50-90 C). I'd like one small enough to } fit into our fume cabinet, but the smallest one I can find } is about 13 x 13 x15 inches. Does anyone know of a source } for a smaller oven? } } Thanks in advance, } } Doug } ---------------------- } Douglas R. Keene } Associate Investigator } Shriners Hospital Research Facilities } 3101 S.W. Sam Jackson Park Road } Portland, Oregon 97201 } phone: 503-221-3434 } FAX: 503-412-6894 (9-5 PST) } e-mail: DRK-at-shcc.org } } } }
-- Gib Ahlstrand Electron Optical Facility, University of Minnesota, CBS Imaging Center, 35 Snyder Hall, St. Paul, MN. USA. 55108 (612)624-3454 (612)624-1799 FAX, giba-at-puccini.cdl.umn.edu http://www.cbs.umn.edu/ic/
Consider this option to an actual oven: I've used a TEMP-BLOK MODULE HEATER (by Lab-Line, distributed by Scientific Products) for curing resins. Its a small electrical heating device measuring about 5x8x3 inches, has high and low temperature ranges with variable control for each. They have a variety of removable blocks with arrays of wells in them into which you can put BEEM capsules, Eppendorf tubes, gelatin capsules,etc. I stick a thermometer in one of the wells to calibrate the tepmerature settings. Place a tight covering of aluminum foil over the top of the block to better stabilize the temperature inside.
I have bought them used from our University's scientific apparatus shop which trades in used lab gear.
Good luck,
Gib
} } Dear Microscopists, } } I am looking for a small oven in which to polymerize epoxy } embedded samples (50-90 C). I'd like one small enough to } fit into our fume cabinet, but the smallest one I can find } is about 13 x 13 x15 inches. Does anyone know of a source } for a smaller oven? } } Thanks in advance, } } Doug} } ---------------------- } Douglas R. Keene } Associate Investigator } Shriners Hospital Research Facilities } 3101 S.W. Sam Jackson Park Road } Portland, Oregon 97201 } phone: 503-221-3434 } FAX: 503-412-6894 (9-5 PST) } e-mail: DRK-at-shcc.org -- Gib Ahlstrand Electron Optical Facility, University of Minnesota, CBS Imaging Center, 35 Snyder Hall, St. Paul, MN. USA. 55108 (612)624-3454 (612)624-1799 FAX, giba-at-puccini.cdl.umn.edu http://www.cbs.umn.edu/ic/
I would like to receive informations about seed tissues separation techniques to analyse under optical microscopy. Thanks for your attention Sinceraly Carla Bocchese
__________________________________________________________________________ Acesso fácil, rápido e ilimitado? Suporte 24hs? R$19,90? Só no AcessoBOL - http://www.bol.com.br/acessobol/
One day while walking down the street a highly successful HR Director was tragically hit by a bus and she died. Her soul arrived up in heaven where she was met at the Pearly Gates by St. Peter himself.
"Welcome to Heaven," said St. Peter. "Before you get settled in though, it seems we have a problem. You see, strangely enough, we've never once had a Human Resources Director make it this far and we're not really sure what to do with you."
"No problem, just let me in," said the woman.
"Well, I'd like to," replied St. Peter, "but I have higher orders. What we're going to do is let you have a day in Hell and a day in Heaven and then you can choose whichever one you want to spend an eternity in."
"Actually, I think I've made up my mind, I prefer to stay in Heaven", said the woman.
"Sorry, we have rules..." And with that St. Peter put the executive in an elevator and it went down-down-down to hell. The doors opened and she found herself stepping out onto the putting green of a beautiful golf course. In the distance was a country club and standing in front of her were all her friends - fellow executives that she had worked with and they were all dressed in evening gowns and cheering for her. They ran up and kissed her on both cheeks and they talked about old times.
They played an excellent round of golf and at night went to the country club where she enjoyed an excellent steak and lobster dinner. She met the Devil who was actually a really nice guy (kinda cute) and she had a great time telling jokes and dancing. She was having such a good time that before she knew it, it was time to leave. Everybody shook her hand and waved goodbye as she got on the elevator. The elevator went up-up-up and opened back up at the Pearly Gates and she found St. Peter waiting for her.
"Now it's time to spend a day in heaven," he said. So she spent the next 24 hours lounging around on clouds and playing the harp and singing. She had a great time and before she knew it her 24 hours were up and St Peter came and got her. "So, you've spent a day in hell and you've spent a day in heaven. Now you must choose your eternity," he said. The woman paused for a second and then replied, "Well, I never thought I'd say this, I mean, Heaven has been really great and all, but I think I had a better time in Hell." So St. Peter escorted her to the elevator and again she went down-down-down back to Hell.
When the doors of the elevator opened she found herself standing in a desolate wasteland covered in garbage and filth. She saw her friends were dressed in rags and were picking up the garbage and putting it in sacks. The Devil came up to her and put his arm around her.
"I don't understand," stammered the woman, "yesterday I was here and there was a golf course and a country club and we ate lobster and we danced and had a great time. Now all there is is a wasteland of garbage and all my friends look miserable."
The Devil looked at her and smiled. "Yesterday we were recruiting you, today you're staff..."
You may want to try dipping just the thermanox portion of your block into liquid nitrogen. The sudden change of temperature will likely loosen the thermanox away from your sample. I do not expect epon to be a problem, but I do know that it works well with Spurrs.
Good luck,
Doug
On Mon, 23 Apr 2001 10:12:10 -0500 "tbargar-at-unmc.edu"-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, } I need advice on embedding Thermanox coverslips. It's } supposed to peel off leaving the monolayer behind, but I'm } not having much luck. I'm using Aralidite 502 as the } embedding medium. Would a another resin work better? I } would appreciate any and all advice. Thanks. } } Tom Bargar } EM Lab } UNMC } 402-559-7347 } tbargar-at-unmc.edu } }
---------------------- Douglas R. Keene Associate Investigator Shriners Hospital Research Facilities 3101 S.W. Sam Jackson Park Road Portland, Oregon 97201 phone: 503-221-3434 FAX: 503-412-6894 (9-5 PST) e-mail: DRK-at-shcc.org
I need to find a commercial source for a calibrated search coil to check stray fields in microscope rooms. Does anyone have any experience with purchasing this item?
John C. Wheatley Lab Manager Arizona State University Center for Solid State Science PSA-213 BOX 871704 Tempe, AZ 85287-1704
You can buy calibrated digital magnetic field meters from F.W. Bell. The are good down to ~ 0.1 mGauss traceable to NIST. I have the Model 4080 which is a triaxial field measurement device.
http://www.fwbell.com
For Acoustic Measurements I use the EXTECH 407727 Digital Sound meter
http://www.extech.com
Nestor
Your Friendly Neighborhood SysOp
=============== } } I need to find a commercial source for a calibrated search coil to check } stray fields in microscope rooms. Does anyone have any experience with } purchasing this item? } } John C. Wheatley } Lab Manager } Arizona State University -- =========================================== Dr. Nestor J. Zaluzec Materials Science Division Building 212 Argonne National Lab 9700 S. Cass Ave Argonne, Illinois 60439 USA Tel: 630-252-7901, Fax: 630-252-4289 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 need to find a commercial source for a calibrated search coil to check stray fields in microscope rooms. Does anyone have any experience with purchasing this item?
John C. Wheatley Lab Manager Arizona State University Center for Solid State Science PSA-213 BOX 871704 Tempe, AZ 85287-1704
Hi John The best bet is to contact your local RS electronics supplier ( find them on http://www.rs-components.com/ )and ask for the ELF Magnetic Field Strength meter part number 212 837. We use it extensively for site tests and have found it to be just as accurate as using a professional test kit. The only difference between this unit and a search coil to a scope is that this device simply tells you if you have a field problem between 20 to 1200Hz where a more expensive search coil will tell you exactly what frequency it is that is causing the field.
Good Luck Luc Harmsen Anaspec, South Africa Technical support on microscopy. Tel + 27 (0) 11 476 3455 Fax + 27 (0) 11 476 7290 anaspec-at-icon.co.za www.anaspec.co.za
Remember, ICEM 15 will be in 2002, Durban, South Africa. www.icem15.com
-----Original Message----- } From: John C. Wheatley [mailto:John.Wheatley-at-asu.edu] Sent: 25 April 2001 01:10 To: Microscopy-at-sparc5.microscopy.com
I need to find a commercial source for a calibrated search coil to check stray fields in microscope rooms. Does anyone have any experience with purchasing this item?
John C. Wheatley Lab Manager Arizona State University Center for Solid State Science PSA-213 BOX 871704 Tempe, AZ 85287-1704
Here, finally, are the responses I got to the query below:
A colleague has asked for recommendations for setting up a digital darkroom (fun to spend someone else's money!). This person would benefit from a really good scanner that could deal with prints, large format negatives (4"x5", 3.25"x4") as well as 35 mm slides. At one time I looked into an Agfa Duoscan T2500. Do any of you have an opinion about this or other suitable scanners? ########## ********** We have the Duoscan 1200 but the 2500 is also a very nice unit--additional (real) resolution and a high O.D. range plus 14 or 16 bit image depth. The Agfa units also come with a built-in transparency plate (rather than having to add on a separate transparency adapter). I find that color fidelity is very good with the Duoscan, and Agfa provides both reflective and transparency calibration standards. I am currently scanning in 3x4 TEM negatives at 12 bits (yield is about 26MB per image), and scan time is fairly rapid. There is one option that you might want to consider: the DIImage unit is made for up to 4x5 negatives, and I think (can't remember the last time I read the specs--the neurons aren't firing today) that resolution is in the 2700 dpi range--even for the 4x5 size. ********** I have an Agfa duoscan, not the 2500 but a duoscan and I love it. I scan everything gels, ex-rays, 2x2,line art and em negatives. The scanner is very versatile. I would recommend it. ********** Got one, love it, wouldn't trade it for the world. OF course, we haven't had it long enough for little nitpicky things to start bugging me, but wow, does it do a nice job. We've scanned Polaroid Type 55 negatives, TEM negatives, TEM prints, Type 52 prints, and AFM 3-D presentations with excellent results. We have even scanned in old 35 mm slides (BW and color) and made prints from them, without enhancement, that are just as pleasing as the original slides. We don't regret the purchase. We were able to work through a local photography supply house and obtain a reconditioned one for a reasonable cost. Our biggest problem right now is finding a printer that will do it justice without mortgaging the farm. ********** } From a vendor: We have sold a large number of the Agfa T2500 into this market with great success.
The T2500 is an excellent scanner for TEM negs. The 2500x2500 optical resolution is high enough to capture fine details and still allow cropping or magnification of small areas of the negative. A 3.5 Dmax will capture details in the dense areas of negatives, slides or prints.
The Duoscan uses what Agfa calls "TwinPlate" design. Unlike most flatbed scanners which scan both reflective and film originals through the glass bed, the Duoscan holds films in a drawer, similar to a glassless negative carrier in an enlarger. No glass means no dust or scratches and no Newton Rings.
The Agfa Fotolook software driver is also excellent. Setting up scan parameters is easy and developing custom film terms is straightforward. You have control over almost every aspect of scanning, including a wide range of Gamma control which is beneficial for scanning very low or high contrast TEM negs.
For scanning 35mm slides, a batch holder accommodates up to 20 mounted 35mm slides. Holders are included for 35mm strips, mounted slides, 120/220 films, 4x5 films and a glass plate for odd size or other transparent originals. I like to scan TEM negs using the 4x5 holder, they fit across the opening and it works very well. ******************* I just bought an Epson Perfection 1240U for home use. I am impressed, So was a professional photographer friend - he was the lab photographer until made redundant recently.
It has a removable transmitted light box/lid combo, just lift off the normal lid from its extending hinge slots (to accommodate thickish documents/thin books?) and fit into the space on the flatbed. It has a separate light switch which normally cuts off when you finish a job - it seems.
It comes with a set of (thin, therefore, flimsy) plastic holders for 4 x 5, 120 roll film plus 35 mm and APS (a double strip holder). 35 mm slides have to be put directly on the flatbed.
It came with free Photoshop LE. "LE" meant no "channels" dialogue box! ******************** We have digital imaging equipment set up to produce photographic quality prints from transmission electron microscope negatives 6.5cm x 9.0cm, reproduce photograph prints for posters, prints from slides and of course convert all types of images suitable for email, to name but a few. The flat bed scanner is:- AGFA ARCUS II Slide scanner:- NIKON Coolscan II
Both have given good service. The quality produced by the ARCUS has not been fully exploited as we feel a compromise between file size and quality has to be a consideration. The software we use is Adobe Photoshop. Our equipment is certainly out of date now, but will be interested to hear what the new machines perform like. ********** There is a simple rule of thumb I use.
Nominal grain size of film is about 10 microns (varies with film speed etc but this is the right order of magnitude). Thus to digitize the film to it's nominal limits your scanner should be able to digitize to better than this spatial dimension.
A simple back of the envelope calculation says a spatial resolution of 10 microns is 2540 - dpi..... and as we all know that must be the optical resolution of the scanner not the interpolated resolution. Scanners at this end are obviously more than you need to digitize photo's and get expensive quickly. Also when you see 2 numbers listed as the scanners resolution, believe only the first number, that is the CCD resolution.
Now add your bit depth. 12 bits is the minimum I would shoot for grayscale image, but if your attempting diffraction work the higher the better (i.e. 14 -16 bits+). For color work obviously multiple the bit depth by 3 one for each primary color (RGB). I've seen a number of 36 bit color scanners but not too many 48 bit ones at } 2540 dpi.
Lastly, bit depth is irrelevant if you don't have a high optical density capabilities otherwise your just digitizing noise. The highest value I believe is an OD of 4.0 but this is for DRUM scanners. Flatbed scanners typically run as low as 2.8, upwards to about 3.4 for the best I've seen in a flatbed. ********** We have a Duoscan T2500, and I really like the resolution we can achieve when scanning any transparency media. There is no holder specifically designed for EM negs, but they fit sideways into the 4x5" holders. It's great for scanning Kodachromes; I was given a slide with a photo of someone (very small image) and was able to scan it in, cropped, at 4000 dpi, and turn that file into a 5x7" print without pixellation. Not bad.
The only drawback is that it can be painfully slow when calibrating. Still, I recommend it as a good, medium-to-high-end film scanner. It's also an excellent flatbed scanner, but with the low-end units available today, it's overkill. ********** Have your colleague check out the Imacon Flextight Precision II scanner. The optical resolution is 5760 dpi for slide-sized objects; I believe it drops to 4800 dpi for objects the size of her larger negatives. The scanner collects 14 bits of usable data per channel, which can be exported as a two bytes per channel, and has a dynamic range of 3.9 OD units (4.1 OD max). The machine is also very fast. The URL is: http://www.imacon.dk/usr/imacon/wppImacon.nsf/pages/flexprecision.html ********** If you are scanning EM negatives, you need to keep the dynamic range in mind. Regular flat beds are closer to 3.2 to 3.4 usually.
The ArtixScan 1100 has a Dmax of 3.9 (about $1600). This was has a 1000 x 2000 dpi resolution. more details at www.microtek.com
The Agfa DuoScan HiD (about $2400) has a 3.7 dynamic range. more details at www.agfa.com
Nikon has the new CoolScan 8000 that has a 4 or 4.2 dynamic range but it doesn't hold the large EM negative size - I think it is limited to something like 2.5 x 3.5 but their website http://www.klt.co.jp/Nikon/Press_Release/ls-8000_main.html has the details.
I think I am going to go with the ArtixScan and buy an extra template and have it machined to hold my size of negatives. Somewhere I saw scary data showing that it is important to support all 4 edges of the negative or you get significantly less optimal scans. The ArtixScan comes with 4 holders but none match my negative size exactly. It has a glass plate holder but the problem with these and any conventional flat bed scanner is that you get Newton rings on many or all of the scans if you look closely.
If you are willing to spend $14,000, there is a really neat film scanner called the Imacon Flextight Precision II CCD Drum Scanner that goes up to 5600 dpi (true optical) and 4.1 Dmax. I wish I could afford it. One web site with info about it is http://www.medgraphix.com/imaconscan.htm
a web site with really strong views on scanning negatives is http://www.flatbed-scanner-review.org/ ********** I have the HP Photosmart film scanner. It has a scanner of 2400 dpi, for 35 mm film. I think the recommendation of a film scanner is a good one for the following reason. Some scanner manufacturers make transparency (slide/negative) devices that use mirrors, but the image quality is poor. The Dimage or other large format scanners should provide acceptable images. The catch for large images and high resolution you need a lot of RAM memory. ********** I have the Duoscan and a Nikon slide scanner. The Duoscan can scan slides on the special tray feature but side by side comparisons of the Duoscan and Nikon show that the Nikon scan is much better. For the larger negs we had a special tray made for the Duoscan and we scan in our EM negs. The Nikon has gotten much cheaper and an excellent scanner can be had for $700 with Digital ICE, something you want. Get two scanners. ********** I love my Epson 1640 scanner, 1600x3200 and up to 4x5 negs and transparencies. ********** I was forwarded your inquiry into digital darkrooms by a colleague. I tackled this issue a few years back and the solution I arrived at is working out fine. I have been a professional photographer for 12 years. I work as an imaging specialist/photographer at a Materials Technology Laboratory. When our lab went digital (not yet 100%), I purchased what was then a very good flatbed scanner - Agfa Arcus II. It was a compromise of sorts. It could handle reflective and transparent originals. It has a max density of 3.2 and a max optical resolution of 1200 dpi. It is fine for in house publications and reports but falls short for anything going to a service bureau. I also don't recommend it for 35mm film. It can scan 35mm but not to the quality I required. We still use the Agfa for many scanning tasks but I have since purchased a more capable machine.
The new scanner is a Flextight Precision II, made by Imacon. It has a Dmax of 4.1, a true optical resolution of 5760 dpi and scans at 14 bits per colour. I purchased it primarily for it's density range. We have a large characterization section with a variety of beam instruments but the TEM negs were always tough to print. Some diffraction patterns take hours to print in a wet darkroom. I used the TEM negs as test samples for the scanners I was considering. A weak point of almost all the prospective film scanners was no holders for TEM film. Imacon has the capability of accepting custom made holders (Imacon will make them based on client specs). As well, the Precision II is primarily a film scanner. It will scan reflective originals up to A4, but I rarely use it for that.
If your colleague is looking for a flat bed scanner, Imacon makes a model called the Progression. It is equally as capable as the Precision but appears to handle reflective originals easier( it accepts film originals from 35mm to 5"x7").It also has a Dmax of 4.1, a true optical resolution of 5760 dpi and scans at 14 bits per colour. These are both quite a step up from the T2500. The 2500 boasts a resolution of 5000 dpi but that's interpolated resolution. I make it a practice not to interpolate when scanning scientific images because of the addition of false image information. The 2500 has a Dmax of 3.4 which is quite acceptable for correctly exposed film or originals with slight underexposure. I don't think it could handle a "Hail Mary" type of neg. With the Precision II, I've pulled quality information off a TEM neg in regions where it seemed transparent to the naked eye. I am very impressed with this machine. I don't want to seem indifferent to the T2500 however. I believe it is a good scanner and can handle most jobs with ease. I would also consider the acquisition software. Fotolook is quite good. I like it's tone curve editor. But Colorflex packaged with the Imacon scanners allows more manual control. It has Photoshop-like unsharp mask controls, good colour correction in all channels, ICC profiles, dot gain compensation etc.
I don't know your colleague's requirements. If he/she is looking for a capable, affordable desktop flatbed, I think you were quite correct to recommend the T2500. If he/she is hoping for more capability I would suggest they look into the Imacon line (www.imacon-usa.com).
The Imacon scanners are comparatively affordable. The Precision II is ~ $14,995 US and the Progression is ~ $19,995 US. I say comparatively because many comparable scanners are much more expensive ( priced between $14,000-$150,000). I realize it is a big jump from the $4500 from the T2500. I justified the expense with not only the quality increase but the time saved in the darkroom with trouble negatives. ********** I used Agfa DuoScan HiD earlier and I try to get it here as well. I like that machine a lot. It's optical resolution is 1000x2000 Dynamic range is 3.7D, which would help scanning DP's. If you want more info you can have a look at: http://www.agfa.com/scanners/duoscan_HiD.html Printing is another task you can buy things from AGFA as well. Their photoprinter is just excellent, but a bit expensive. I have tried nice HP inkjet printers with great success. ********** In response to Tina's post, I have not seen any mention on the list of the scanner I purchased a few weeks ago, the Epson Expression 1640XL. It has 1600dpi optical resolution (scans at a hardware resolution of 1600x3200 dpi) 42 bit color (14 bit gray) and Dmax of 3.6. It is large format, and the transparency adapter comes with a range of negative holders. Has SCSI or USB interfaces with firewire as an optional extra (I use USB on a Win 2000 system). Of course, you pay for what you get - it isn't cheap.
We are only just beginning to learn how best to use all the resolution and bit depth we now have, but I and my users love it!
This is not a comparison, of course (I haven't used the other models) but just to say we are happy with what we have. ********** We are getting first rate resolution results from our "UMAX Powerlock 1100 Magicscan" scanner coupled to a" FUJIX Pictography 3000" printer. Our base computer is always an Apple system upgraded periodically. ********** } In response to Tina's post, I have not seen any } mention on the list of the scanner I purchased } a few weeks ago, the Epson Expression 1640XL. } It has 1600dpi optical resolution (scans at } a hardware resolution of 1600x3200 dpi) 42 bit } color (14 bit gray) and Dmax of 3.6.
I would certainly believe the resolution and the color depth for this scanner is adequate, but if scanning TEM films is an issue, I'd seriously advise measuring the optical density of your films ... I've heard these approach OD} 4 ... which would imply you might consider the dedicated film scanners, e.g., Polaroid 45 Ultra or the new Nikon LS-8000. ********** I have the scanner you are looking at & like it a lot. To be quite honest I do not find that I need to exploit it's full capabilities. If I were in the market again, looking at newer technology I would be interested in a faster scanner of similar quality. Yes I want my cake & to eat it too :). I'll give you this analogy. If I have 10 negatives I will franchise my time, that is let things scan while I hang out in the office doing other things. If I have 20 negatives, I'll probably go to the darkroom to make photos. It is quicker & paper is cheaper. BTW I have an Epson 870 inkjet that produces nice quality images... cost is down to $180 US, (now the Epson 880)....no financial interest in these companies. ********** There was a thread recently on scanners for TEM film. I have looked up all the models mentioned, on the web and called agents for prices - and produced a comparative table, given below. I do not guarantee that the figures are accurate but they are my best interpretation of the data given. In the light of experience and Nestor's comments, I would suggest that 2000 dpi is a minimum for TEM negatives. You may be able to get away with less nine times out of ten, but there will be occasions when you need more. I would exclude the Minolta and all the Epsons from consideration (despite the incredibly low prices of some of the Epsons) because of the low pixel density. Among the rest the Nikon has the best pixel density and the best optical density (another critical parameter for TEM negatives). The price is very competitive too. The Nikon web site does not give a time for scanning a negative. On the face of it the Nikon would be a best buy - get a separate, inexpensive flatbed scanner for the other work. These comments are all my own opinions based on manufacturers' data. Since we are considering purchase any comments to the contrary would be most welcome. Code Maker Model Type A Agfa DuoScan T2500 Flatbed -Transparency included
B Epson 1640 several versions Flatbed -Transparency option 1680 several versions
C 1600 several versions Flatbed -Transparency included
D Imacon Flextight Precision II Drum -for film and large format
E Minolta Dimage ScanMulti II Film
F Nikon Super Coolscan 8000ED Film
G Polaroid 45 Ultra Film
H Umax Powerlook 3000 Flatbed -Transparency included
Code dpi OD Time Price Opinion at 6 x 9 cm
A 2500 x2500 3.4 3 min $4,500 Fair B 1600 x 3200 3.6 $300-$3000 Poor $800-$1400 Poor
C 1600 x 3200 3.3 $650-$1160 Not suitable
D 2240 x2240** 3.9/4.1 N/A above $10k Good: low pixel density
E 1128 x 1128 3.6 Not suitable
F 4000 x 4000 4.2 N/A $2,695 V. Good
G 2500 x 2500 3.8 5 min $7,495 Good but pricey
H 3048 x 3048 3.6 3 min $6,499
********** I too am about to buy and I would make a couple of comments on your evaluation. First, let me remind everyone that the Dynamic range is a log scale so small numerical differences are significant.
I also think the Nikon Coolscan 8000 looks great but it only takes a 2.5 x 3.5 negative which is smaller than my JEOL and Hitachi EM negative sizes (~ 3 1/2 by 4 1/2"). Have these EM manufacturers gone to a smaller film size or is Nikon using a non-Japanese EM as their standard? seems odd but I don't see how the Nikon would be very useful. You say a {2000 line scanner would be useful 9 out of 10 times but want the 2000+ lines for the occasional high res scan. I would argue that the size of the negative was the more important variable to be worried about. The Nikon couldn't handle 4x5 LM negatives or transparencies from autoradiography of Westerns/Northerns, etc.
My leading candidate is the ArtixScan 1100 has a Dmax of 3.9 (about $1600 with SCSI card). This was has a 1000 x 2000 dpi resolution. more details at www.microtek.com. This is my leading candidate. It was 4 negative carriers and I await word whether one could be modified to carry a 3 1/2 by 4 1/2 negative. At worst, I will have my scientific instrumentation shop guys fabricate a holder. It comes with a glass 8 x 10 glass carrier for odd size negs but I want to avoid Newton rings and want a glassless carrier.
I would appreciate comments on the following argument (I think I have this correctly figured out but am not sure since so many out there seem to want to have a higher resolution scanner). I have a Fuji Pictrography 3000 printer with a 400 dpi output that is as good as any other widely available printer in the academic world. If you figure the maximum published image size is about 8 inches, that would mean the maximum image size be 3200 dpi wide. A 1000 dpi scan of my negative would be 4500 x 3500 dpi. I could crop by about 28% or 10% depending on the orientation of the negative and still be taking full advantage of the printer resolution. In reality, most EM publication prints are smaller than 8" wide so one could crop even more and still not need more than 1000 dpi. A resolution } 1000 dpi would be useful for subtle morphometric analysis but a 4000 dpi scan of a 3 x 4 negative would be 192 MB. That is pretty big for doing morphometry on! A 1000 dpi scan of a 3.5 x 4.5" negative would be about 16 MB and that is much more manageable. Perhaps the difference is in the type of EM we are doing. I am working with biological specimens doing standard thin section type stuff. are you doing some Material Sci application that demands more? I would love to take advantage of the Firewire option but my information is that the 8700 has a Dmax of 3.4 vs the 3.9 for the 1100. That is a significant difference. Do EM negatives of biological thin sections reach that? I think so. I do a lot of EM immunocytochemistry and have to look for gold (intensely black) against a very dark tissue component so I am hoping the higher Dmax improves my results. I frequently scan negatives on a Umax 1100 (Dmax 3.4??) and can't differentiate the gold from the background although by eye I can discriminate them when the negative is placed on a light box. Changing my exposure would give me an unusable image for the rest of the tissue. Maybe this is an extreme case but I suspect that lots of "dark organelles" (e.g., lysosomes, nuclei) have fine structure that get lost in the scanning with a low Dmax scanner. ********** Your information is correct and mine is not. The Dmax of the 8700 is 3.4. ********** A colleague and I each recently bought Microtek scanners to scan TEM negatives. I have the Artixscan 1100 and he has the Model 8700 which has similar characteristics (actually higher resolution -1200dpi), 3.9 dmax at 42 bits color (14 grayscale), and the glassless film carrier setup. The 8700 has USB and Firewire interfaces and is cheaper ( {$1000), and the 1000 dpi Model 1100 has a SCSI interface. You might want to check out the specs of the lower cost model 8700 on the microtekusa website if your computer can handle USB or Firewire. Both scanners have performed up to our expectations, which I would characterize as modest. Microtek does not supply a 3-1/4 x 4 " negative carrier for standard size TEM film but you can easily make a serviceable one from stiff paper or light cardboard.
How much scanner resolution should you buy? The answer depends on how you intend to use it. Most applications do not require capturing the full resolution of the negative. From a practical viewpoint, the scanner resolution just determines how many times you can magnify the negative image to produce the final print size. For example, to get a publication-size print at 300 dpi, an image scanned at 1200 dpi scan could be zoomed 4X. A practical alternative to spending more for higher scanning resolution is to take photos at higher magnification. One exception is with lattice images from the TEM, which (depending on the lattice fringe spacing on the negative) might require higher scan resolutions to avoid getting a moire effect. (Of course, not everyone agrees. My colleague prefers to always scan at the maximum resolution).
What does a Dmax of 3.9 mean to you? To me it means a very dark negative. D is the log of the transmitted to incident intensity ratio. I wonder if users ever actually verify the manufacturer's specs with a calibrated density target. A Dmax of 3.9 can be useful for scanning TEM diffraction patterns that might have high contrast, but TEM micrograph negatives of metals and ceramics generally don't have that much contrast and biological thin section photos tend to have rather weak contrast. If your negatives are simply dark, use shorter photo exposure times. Scanning with maximum allowed grayscale resolutions (e.g., 14 bits rather than 8) is highly recommended if you intend to enhance or adjust images, but that's another story. I believe that those Agfa scanners are OEM by Microtek. If budget is the concern, I would recommend buying a Microtek Scanmaker 5 ($1,100) instead, which I have used for scanning quite a few EM negatives and have satisfactory results. The Dmax and the dynamic range for Scanmaker 5 is are about 3.7 and 3.4. Another model your colleague might consider is an Agfa Duoscan HiD ($ 2,500) which has a higher Dmax of 3.7, but less optical resolution of 1,000 dpi compared with 2500 dpi on a DuoScan T2500. What others failed to mentioned is that the DuoScan T2500 only has a narrow strip on the CCD bay being capable of scanning at 2,500 dpi, otherwise the true optical resolution is 1,000 dpi. Although a lot of investigators think that the higher scanning resolution, the better, my personal bias is leaning toward to purchase a scanner having at optical resolution at 1,000-1,200 dpi. Umax also carries a few mid- to high-end scanners such as Powerlook III for routine negative stains. My personal experience for the UMAX scanner is only limited to the Powerlook II, a mid-range scanner which gives more grayish scanned images compared to those high end models I mentioned previously. However, it is a descent scanner if you are working with color transparencies.
********** I have not summarized the lengthy thread about film (logarithmic) vs digital (linear) response!
Aloha, Tina
************************************************* * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
} I am helping a former traditional photographic media user/computerphobe } set up digital imaging capabilities since her university/museum } department is closing their darkroom facilities and reassigning personnel } (sigh). She has what appears to be a decent budget (until I started } pricing the good stuff!). I am proposing she get a fast (733MHz) G4 Mac } with maximum (1.5GB) RAM, and she saw and fell in love with the Apple } 22" cinema display (as did I when I saw it in person). People seem to } like the Agfa T2500 scanner for prints and negatives. A moderate color } printer,since she has access to other really good printers in her department. A } Nikon Coolpix 990 digital camera for on- and off-microscope. Photoshop } 6.0, for which I'll train her. Corel Draw for vector graphics? ********** Adobe Illustrator or Freehand for vector graphics. Corel Draw is less common and has a proprietary file format that has caused me troubles preparing articles for Microscopy Today. Corel can save in other formats, but people have to use that option. Also, Adobe has educational pricing, and special package prices that bundle full versions of Photoshop and Illustrator.
The 733 Mhz G4 Mac is a great idea since it comes with the superdrive for making DVDs and CDs, which your friend will probably find very helpful for archiving. I would also encourage getting a copy of Retrospect for backups. I am a huge fan of HP printers and prefer them over Epson. I find the print quality as good or a bit better, and the software engines are much faster, in my experience with HP 970 cxi and Epson 750 and 850.
I also very much like the Nikon 990, although it has taken me a long time to figure out the best way to use it (almost too many options....especially for white balance and light metering, but the quality is very good). Make sure she gets a memory disk for the 990 that is large enough, 64 MB at least. If the memory card is too small she won't be able to save TIFF files, only JPG. She will need TIFF if anyone wants to do any sort of quantitative imaging, and the TIFF files are all well over 2 MB each. Get at least two cards and a USB card reader (these are cheap) so she can keep the card reader attached to the Mac, and just swap out memory cards instead of connecting the camera directly to the Mac, which is kind of a pain. Make sure she gets the AC adapter, too. Running the Nikon 990 off batteries is OK if you need to be mobile but in a lab setting swapping batteries gets old. Alternately, get a battery charger and extra NiCad batteries and keep the battery charger going all the time in a handy outlet. Then you avoid the extra AC power cord, which reduces cord tangle (a pet peeve of mine....). ********** If you want to do a darkroom for $10K you can do pretty well, especially if you don't need to include a killer printer. If you need a decent printer for proofing, etc, I would either use an inkjet or a low/mid end laser printer. Does she need color output from the local printer?
For the scanner, if she wanted one "great" scanner, the T2500 would be my choice, but here's a few things to consider. What makes a scanner great depends on the workflow.She should look at her workflow, in particular what amount of scanning from 35mm will be done? If there is much 35mm, will there be a large amount of 35mm slides where some type of auto feeder would be handy? Would scanning from uncut rolls of 35mm film be needed? The mix is the first thing to figure out. For occasional 35mm scanning the T2500 would do the job. As the mix of 35mm increases, the value of a dedicated 35mm scanner increases. One of the best new 35mm scanners will be the Nikon Supercoolscan 4000ED. 4000dpi res, great Dmax, plus a slide feeder, long roll holder and even an adapter for glass microscope slides are available.
On the TEM side, I think the T2500 is still the nuts, especially for cropping a smaller areas of a neg and still having the resolution to blow it up for larger prints.
If the scanner budget didn't allow for the T2500 or for the T2500 along with a 35mm scanner, the Duoscan Hi-D would be my next choice. The Hi-D has slightly lower res(2000x1000) but a high Dmax of 3.7. Agfa's software is also very powerful.
The first key is probably to figure how much she's going to spend on the workstation. After that do the printer and/ or the scanner(s).
Microtek has a new line called Artixscan. When you look at the specs you will notice many similarities to the Duoscans. This is because Microtek shares some of Agfa's hardware. Agfa makes their own apochromatic lenses and the software is different. I personally like Agfa's software better but Microtek does offer a slight price advantage. Attached is info on the Microtek 2500 and 1100 scanners. ************************* I both run the microscope labs here and am a researcher in materials engineering.
First I want to say that I strongly support the use of the digital laboratory. While we still occasionally use film for our highest quality requirements, in general we are fully digital. The use of digital cameras has really expanded our undergraduate teaching laboratories and has sped up our research.
I have found one "dark-side" to a digital imaging laboratory as a lab manager. As the lab manager, I have found that keeping a digital laboratory up-to-date is much more expensive than the film laboratory. When we were only film, we had to repair the film cartridges for our Polaroid PN film (it takes about five minutes) and have the microscopes cleaned about once a year at a cost of about $1k.
The digital lab. is much more expensive time and repair wise. Because we crunch our computers with our image size and storage, it takes more of my time to keep stuff going. All our computers are networked and in addition to work, the students tend to junk up the computers with downloads etc which stops them from working for the image processing work. This requires continual monitoring on my part (in spite of rules against using them for these applications!) In addition, keeping computers that will run the data is expensive. I buy pretty much the best out there, but somehow upgrades are still inevitable. I also have to supply print cartridges, etc. Researchers always supplied their own film and dark room supplies. In addition, I've had to have our cameras repaired numerous times. The cost was high (at least $500) and they stayed gone for up to a month. Finally, some of my cameras are about 3 years old. I can see a degradation in the image quality from when they were purchased. The cameras are much noisier. I see a future of regular replacement of my cameras in addition to the computer upgrades. So while the cost to the researchers is lower (which helps me as a researcher), the cost to the lab itself is higher (which hurts me as a lab manager). I'm working on setting up a fee schedule for this equipment but REALLY hate to have to do it. All of you who do this in a university know how painful it is!
Regarding the camera purchase-in addition to considering the camera resolution and cost, I think you should consider the image transfer. I recommend considering a camera with immediate transfer of the image to the microscope if you have numerous inexperienced users. Being able to focus on the screen is extremely helpful. The image transfer time is also important if you have many images to capture. We do image analysis on numerous images and some of the cameras have about a 30 second transfer time for decent resolution. This would be unbearable for the number of images we collect. I'm not sure how the Nikon Coolpix works but this should be considered by anyone that is purchasing a digital camera. ********** I just put together a nearly identical system: G4 533MHz, 1.5GB, 22" conventional monitor (but loved the cinema), Agfa and Polaroid 4x5 scanners, Nikon 990 and Fuji Pro S1, Photoshop, Adobe Illustrator, HP 5000PS poster printer and Adobe In-Design for laying out posters. Whew.....awesome.
Glad to see that you went with Macintosh. Very wise. Too many people fall for the empty promises made by the PC Windoze. ********** I agree that Corel Draw is less common, *especially for Macintosh*. If you were setting up a PC studio, I'd say otherwise, but Illustrator and Freehand have been the standard tools for Mac vector graphics for about a decade. I think Illustrator may have expanded portability with Photoshop since they're both made by Adobe. For the most part, Illustrator, Freehand and CorelDraw provide more or less the same features, they may just call their tools by different names. Once you know one, it's not too difficult to pick up the others for basic illustration. ********** I think you are on the right track. I think you might look at a dedicated slide scanner too. We got one years ago and it has seen lots of use. So many people have big collections of slides that they want to turn into digital images and the slide scanner fills the gap. It is quick, easy, and doesn't need much training to use. We also have a flat bed scanner, Arcus II, that gets lots of use, but I am glad we have the slide scanner too. Arranging slides and cropping etc, can be a pain on the flat bed. With the slide scanner, just slip in the slide and scan away. Ours is an old Polaroid Sprint Scan. Today you can get one for pretty cheap that is even better than ours.
I agree with the recommendation to go with Illustrator over Corel. Illustrator plays well with Photoshop and has never screwed us up. Sometimes we run into problems, Canvas has also been a trouble maker.
After a while we started having 'Disk Full' errors on the machine used with the scanners. Photoshop wants at least 2 - 3 times the size of the file on the 'Scratch Disk' to swap files etc. If the disk is getting full, you get stopped by no room on the 'Scratch Disk', same can happen when trying to print big files, need room to spool the file for printing. So, get some kind of removable medium, new Macs might have a CD-RW and that would be cool. I partitioned our big drive, setting aside 1 GB as a scratch disk where no files or other junk are allowed.
Slightly off the main topic, we have found a wide format printer gets lots of use. We have an HP 755CM. 36" wide color inkjet. People from all over use it to make posters for meetings and displays for classes etc.
I think you have started a very good discussion. It is one all of us are facing and things are changing so fast, we all need to benefit from the experience of each other. I don't mind at all that you 'introduced a subject that gets periodically posted here', its a new ballgame just about every six months. ********** } From last February(?)
} A broad question for the light microscopists- } } I'm writing up a wish list for our EM lab, and it includes (gasp) light } microscopes. My question is - how do I go about evaluating and choosing a } digital camera for light microscopes? It would be for both compound and } dissecting microscopes, should be color, decent resolution, not } necessarily low light nor real-time video, but capable of good images for } image analysis on sections. We are getting a confocal, so fluorescence } imaging would be done there rather than with the proposed 'scope and } camera. } } What do I need to look for, and what price ranges are we talking about? ********** Here is what I know from my explorations. We got a Kodak MDS 120 system. It is now obsolete and has been replace by a newer, higher resolution model. Cost was a couple of K. Half the cost was the adapter to get the camera on to the microscope.
Unlike many 35mm cameras, the lens of most digital cameras is not removable. So to get the camera to see what the microscope sees, there needs to be this special adapter gizmo. It is matched to the threads on the camera at one end and then to the microscope on the other end. The DC 120 is about 1K x 1K res. and does pretty well for up to 5" x 7" pictures. Even some 8" x 10" are OK. The key is to remember is a photo 'documentation' system. Good for reminding yourself of what you saw and good enough for most applications, but not the equal of film.
Using it is similar to film, only different. It works best in brightfield with plenty of light. It is rated at an equivalent of ASA 160. The overall appearance of the pictures looks harsher to me. The dynamic range seems to be more compressed than film. Adjusting the light for best contract is different than using film.
The biggest negative is the way it transfers the digital file from the camera to the computer. It goes something like this: Set up the camera like a film camera, fiddle around with the adapter (it is focusable) until the eyepieces and the camera are parfocal (not easy). If you are pretty close, take a preview picture. Wait. Wait. The picture has to be transferred from the camera to the computer. It is not real time like a video camera. After you wait, the image appears on the computer screen. If its OK, usually not, take a final high resolution picture. Wait longer while the larger file is transferred. Once its there, you are done. Ours uses a Photoshop plug in for acquisition so it easy going from here.
The trouble is that it is more like film in that you don't know if the picture is good the instant you expose it. Sometimes the light is not just right, sometimes the focus is a little off. You can't see these things in real time like you can with a video camera, but a video camera is really crummy resolution. If everything is set perfectly, then it is like exposing film, you take the picture and wait, a few seconds to a minute rather than a few days to a week to see the final result. I never worried about how the pictures would come out with film, they always did. But somehow not being able to see the final result instantly with the digital camera is very frustrating. Maybe since the digital camera is an add on it is not as well set up and out film camera for focus etc. I did find out that I had to get some non-adjustable eyepieces so the focus between the oculars and the camera would not change between users. We went round and round chasing focus until that was fixed.
This is another problem if you plan to use the camera on different microscopes. The adapter will have to be adjusted to the new system each time it is changed. A pain.
All that said, it has been useful having the camera. I learned a lot about digital photography, some people have gotten good use from it on a microscope, it is useful for regular photography, and I now can sound like an expert when discussing the pros and cons of systems with folks here and abroad who want to buy a system. ********** I looked at the MDS 290 also, Kodak was offering an 'upgrade' path from the 120. I passed because the 290's equiv. ASA is 100 vs 160 for the 120. Also the 290 would require USB for image transfer. It is supposed to be faster, but still won't be video rate. Also to get USB in the room with the microscope would require a new computer and since the old computer has a NuBus frame grabber for our video camera I would have to get a new frame grabber for the PCI bus new computer. So while the camera upgrade was pretty attractive, the total cost to upgrade was way more than could be justified by the amount of use that the digital camera has generated so far. I have heard a lot of interest in the Nikon Cool Pix 990 (I think that's the one). It apparently has the ability to send a lower res. B&W signal to a video monitor so focusing etc can be done in real time, then switch to high res color for the final digital shot. ******************************
Thanks to everyone who contributed!
Aloha, Tina **************************************************************************** * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * * Biological Electron Microscope Facility * (808) 956-6251 * * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* ****************************************************************************
I wondered how commonly people use a tool (like a loop) to pick up ultrathin sections.
Does it make life easier or does it only introduce different problems?
I was told that I can buy some "luxury" - but am still undecided.
Your views would be very much appreciated.
Regards
Claudia
Dr. C. Hayward-Costa School of Life Sciences Kingston University Penrhyn Road, Kingston upon Thames Surrey KT1 2EE, UK 44(0)208 547 2000 x 2240 Email: c.hayward-at-kingston.ac.uk Fax: 44(0)208 547 7562
Tina: Thanks for the time and effort you have invested in this. A most enlightening and useful summary.
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc On Tue, 24 Apr 2001 21:58:02 -1000 (HST), Tina Carvalho 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, finally, are the responses I got to the query below: | | A colleague has asked for recommendations for setting up a digital | darkroom (fun to spend someone else's money!). This person would benefit | from a really good scanner that could deal with prints, large format | negatives (4"x5", 3.25"x4") as well as 35 mm slides. At one time I looked | into an Agfa Duoscan T2500. Do any of you have an opinion about this or | other suitable scanners? | ########## | ********** | We have the Duoscan 1200 but the 2500 is also a very nice unit--additional | (real) resolution and a high O.D. range plus 14 or 16 bit image | depth. The Agfa units also come with a built-in transparency plate | (rather than having to add on a separate transparency adapter). I find | that color fidelity is very good with the Duoscan, and Agfa provides both | reflective and transparency calibration standards. I am currently | scanning in 3x4 TEM negatives at 12 bits (yield is about 26MB per image), | and scan time is fairly rapid. There is one option that you might want to | consider: the DIImage unit is made for up to 4x5 negatives, and I think | (can't remember the last time I read the specs--the neurons aren't firing | today) that resolution is in the 2700 dpi range--even for the 4x5 size. | ********** | I have an Agfa duoscan, not the 2500 but a duoscan and I love it. I scan | everything gels, ex-rays, 2x2,line art and em negatives. The scanner is | very versatile. I would recommend it. | ********** | Got one, love it, wouldn't trade it for the world. OF course, we haven't | had it long enough for little nitpicky things to start bugging me, but | wow, does it do a nice job. We've scanned Polaroid Type 55 negatives, TEM | negatives, TEM prints, Type 52 prints, and AFM 3-D presentations with | excellent results. We have even scanned in old 35 mm slides (BW and | color) and made prints from them, without enhancement, that are just as | pleasing as the original slides. We don't regret the purchase. We were | able to work through a local photography supply house and obtain a | reconditioned one for a reasonable cost. Our biggest problem right now is | finding a printer that will do it justice without mortgaging the farm. | ********** | } From a vendor: We have sold a large number of the Agfa T2500 into this | market with great success. | | The T2500 is an excellent scanner for TEM negs. The 2500x2500 optical | resolution is high enough to capture fine details and still allow cropping | or magnification of small areas of the negative. A 3.5 Dmax will capture | details in the dense areas of negatives, slides or prints. | | The Duoscan uses what Agfa calls "TwinPlate" design. Unlike most flatbed | scanners which scan both reflective and film originals through the glass | bed, the Duoscan holds films in a drawer, similar to a glassless negative | carrier in an enlarger. No glass means no dust or scratches and no Newton | Rings. | | The Agfa Fotolook software driver is also excellent. Setting up scan | parameters is easy and developing custom film terms is | straightforward. You have control over almost every aspect of scanning, | including a wide range of Gamma control which is beneficial for scanning | very low or high contrast TEM negs. | | For scanning 35mm slides, a batch holder accommodates up to 20 mounted | 35mm slides. Holders are included for 35mm strips, mounted slides, | 120/220 films, 4x5 films and a glass plate for odd size or other | transparent originals. I like to scan TEM negs using the 4x5 holder, they | fit across the opening and it works very well. | ******************* | I just bought an Epson Perfection 1240U for home use. I am impressed, So | was a professional photographer friend - he was the lab photographer until | made redundant recently. | | It has a removable transmitted light box/lid combo, just lift off the | normal lid from its extending hinge slots (to accommodate thickish | documents/thin books?) and fit into the space on the flatbed. It has a | separate light switch which normally cuts off when you finish a job - it | seems. | | It comes with a set of (thin, therefore, flimsy) plastic holders for 4 x | 5, 120 roll film plus 35 mm and APS (a double strip holder). 35 mm slides | have to be put directly on the flatbed. | | It came with free Photoshop LE. "LE" meant no "channels" dialogue box! | ******************** | We have digital imaging equipment set up to produce photographic quality | prints from transmission electron microscope negatives 6.5cm x 9.0cm, | reproduce photograph prints for posters, prints from slides and of course | convert all types of images suitable for email, to name but a few. | The flat bed scanner is:- AGFA ARCUS II | Slide scanner:- NIKON Coolscan II | | Both have given good service. The quality produced by the ARCUS has not | been fully exploited as we feel a compromise between file size and quality | has to be a consideration. The software we use is Adobe Photoshop. | Our equipment is certainly out of date now, but will be interested to hear | what the new machines perform like. | ********** | There is a simple rule of thumb I use. | | Nominal grain size of film is about 10 microns (varies | with film speed etc but this is the right order of magnitude). | Thus to digitize the film to it's nominal limits your scanner should | be able to digitize to better than this spatial dimension. | | A simple back of the envelope calculation says a spatial resolution | of 10 microns is 2540 - dpi..... and as | we all know that must be the optical resolution of the | scanner not the interpolated resolution. Scanners at this | end are obviously more than you need to digitize photo's and | get expensive quickly. Also when you see 2 numbers listed | as the scanners resolution, believe only the first number, that is | the CCD resolution. | | Now add your bit depth. 12 bits is the minimum I | would shoot for grayscale image, but if your attempting | diffraction work the higher the better (i.e. 14 -16 bits+). | For color work obviously multiple the bit depth by 3 | one for each primary color (RGB). I've seen a number | of 36 bit color scanners but not too many 48 bit ones at | } 2540 dpi. | | Lastly, bit depth is irrelevant if you don't have a high | optical density capabilities otherwise your just digitizing | noise. The highest value I believe is an OD of 4.0 but | this is for DRUM scanners. Flatbed scanners typically | run as low as 2.8, upwards to about 3.4 for the best | I've seen in a flatbed. | ********** | We have a Duoscan T2500, and I really like the resolution we can achieve | when scanning any transparency media. There is no holder specifically | designed for EM negs, but they fit sideways into the 4x5" holders. It's | great for scanning Kodachromes; I was given a slide with a photo of | someone (very small image) and was able to scan it in, cropped, at 4000 | dpi, and turn that file into a 5x7" print without pixellation. Not bad. | | The only drawback is that it can be painfully slow when calibrating. | Still, I recommend it as a good, medium-to-high-end film scanner. It's | also an excellent flatbed scanner, but with the low-end units available | today, it's overkill. | ********** | Have your colleague check out the Imacon Flextight Precision II | scanner. The optical resolution is 5760 dpi for slide-sized objects; | I believe it drops to 4800 dpi for objects the size of her larger | negatives. The scanner collects 14 bits of usable data per channel, | which can be exported as a two bytes per channel, and has a dynamic | range of 3.9 OD units (4.1 OD max). The machine is also very fast. | The URL is: | http://www.imacon.dk/usr/imacon/wppImacon.nsf/pages/flexprecision.html | ********** | If you are scanning EM negatives, you need to keep the dynamic range in | mind. Regular flat beds are closer to 3.2 to 3.4 usually. | | The ArtixScan 1100 has a Dmax of 3.9 (about $1600). This was has a 1000 x | 2000 dpi resolution. more details at www.microtek.com | | The Agfa DuoScan HiD (about $2400) has a 3.7 dynamic range. more details | at www.agfa.com | | Nikon has the new CoolScan 8000 that has a 4 or 4.2 dynamic range but it | doesn't hold the large EM negative size - I think it is limited to | something like 2.5 x 3.5 but their website | http://www.klt.co.jp/Nikon/Press_Release/ls-8000_main.html has the | details. | | I think I am going to go with the ArtixScan and buy an extra template and | have it machined to hold my size of negatives. Somewhere I saw scary data | showing that it is important to support all 4 edges of the negative or you | get significantly less optimal scans. The ArtixScan comes with 4 holders | but none match my negative size exactly. It has a glass plate holder but | the problem with these and any conventional flat bed scanner is that you | get Newton rings on many or all of the scans if you look closely. | | If you are willing to spend $14,000, there is a really neat film scanner | called the Imacon Flextight Precision II CCD Drum Scanner that goes | up to 5600 dpi (true optical) and 4.1 Dmax. I wish I could afford it. | One web site with info about it is | http://www.medgraphix.com/imaconscan.htm | | a web site with really strong views on scanning negatives is | http://www.flatbed-scanner-review.org/ | ********** | I have the HP Photosmart film scanner. It has a scanner of 2400 dpi, for | 35 mm film. I think the recommendation of a film scanner is a good one | for the following reason. Some scanner manufacturers make transparency | (slide/negative) devices that use mirrors, but the image quality is poor. | The Dimage or other large format scanners should provide acceptable | images. The catch for large images and high resolution you need a lot of | RAM memory. | ********** | I have the Duoscan and a Nikon slide scanner. The Duoscan can scan slides | on the special tray feature but side by side comparisons of the Duoscan | and Nikon show that the Nikon scan is much better. For the larger negs we | had a special tray made for the Duoscan and we scan in our EM negs. The | Nikon has gotten much cheaper and an excellent scanner can be had for $700 | with Digital ICE, something you want. Get two scanners. | ********** | I love my Epson 1640 scanner, 1600x3200 and up to 4x5 negs and | transparencies. | ********** | I was forwarded your inquiry into digital darkrooms by a colleague. I | tackled this issue a few years back and the solution I arrived at is | working out fine. I have been a professional photographer for 12 years. I | work as an imaging specialist/photographer at a Materials Technology | Laboratory. | When our lab went digital (not yet 100%), I purchased what was then a | very good flatbed scanner - Agfa Arcus II. It was a compromise of | sorts. It could handle reflective and transparent originals. It has a max | density of 3.2 and a max optical resolution of 1200 dpi. It is fine for in | house publications and reports but falls short for anything going to a | service bureau. I also don't recommend it for 35mm film. It can scan 35mm | but not to the quality I required. We still use the Agfa for many scanning | tasks but I have since purchased a more capable machine. | | The new scanner is a Flextight Precision II, made by Imacon. It has a Dmax | of 4.1, a true optical resolution of 5760 dpi and scans at 14 bits per | colour. I purchased it primarily for it's density range. We have a large | characterization section with a variety of beam instruments but the TEM | negs were always tough to print. Some diffraction patterns take hours to | print in a wet darkroom. I used the TEM negs as test samples for the | scanners I was considering. A weak point of almost all the prospective | film scanners was no holders for TEM film. Imacon has the capability of | accepting custom made holders (Imacon will make them based on client | specs). As well, the Precision II is primarily a film scanner. It will | scan reflective originals up to A4, but I rarely use it for that. | | If your colleague is looking for a flat bed scanner, Imacon makes a model | called the Progression. It is equally as capable as the Precision but | appears to handle reflective originals easier( it accepts film originals | from 35mm to 5"x7").It also has a Dmax of 4.1, a true optical resolution | of 5760 dpi and scans at 14 bits per colour. These are both quite a step | up from the T2500. The 2500 boasts a resolution of 5000 dpi but that's | interpolated resolution. I make it a practice not to interpolate when | scanning scientific images because of the addition of false image | information. The 2500 has a Dmax of 3.4 which is quite acceptable for | correctly exposed film or originals with slight underexposure. I don't | think it could handle a "Hail Mary" type of neg. With the Precision II, | I've pulled quality information off a TEM neg in regions where it seemed | transparent to the naked eye. I am very impressed with this machine. I | don't want to seem indifferent to the T2500 however. I believe it is a | good scanner and can handle most jobs with ease. I would also consider the | acquisition software. Fotolook is quite good. I like it's tone curve | editor. But Colorflex packaged with the Imacon scanners allows more manual | control. It has Photoshop-like unsharp mask controls, good colour | correction in all channels, ICC profiles, dot gain compensation etc. | | I don't know your colleague's requirements. If he/she is looking for a | capable, affordable desktop flatbed, I think you were quite correct to | recommend the T2500. If he/she is hoping for more capability I would | suggest they look into the Imacon line (www.imacon-usa.com). | | The Imacon scanners are comparatively affordable. The Precision II is ~ | $14,995 US and the Progression is ~ $19,995 US. I say comparatively | because many comparable scanners are much more expensive ( priced between | $14,000-$150,000). I realize it is a big jump from the $4500 from the T2500. I | justified the expense with not only the quality increase but the time | saved in the darkroom with trouble negatives. | ********** | I used Agfa DuoScan HiD earlier and I try to get it here as well. I like | that machine a lot. It's optical resolution is 1000x2000 Dynamic range is | 3.7D, which would help scanning DP's. If you want more info you can have a | look at: | http://www.agfa.com/scanners/duoscan_HiD.html | Printing is another task you can buy things from AGFA as well. Their | photoprinter is just excellent, but a bit expensive. I have tried nice HP | inkjet printers with great success. | ********** | In response to Tina's post, I have not seen any mention on the list of the | scanner I purchased a few weeks ago, the Epson Expression 1640XL. It has | 1600dpi optical resolution (scans at a hardware resolution of 1600x3200 | dpi) 42 bit color (14 bit gray) and Dmax of 3.6. It is large format, and | the transparency adapter comes with a range of negative holders. Has SCSI | or USB interfaces with firewire as an optional extra (I use USB on a Win | 2000 system). Of course, you pay for what you get - it isn't cheap. | | We are only just beginning to learn how best to use all the resolution and | bit depth we now have, but I and my users love it! | | This is not a comparison, of course (I haven't used the other models) but | just to say we are happy with what we have. | ********** | We are getting first rate resolution results from our "UMAX Powerlock | 1100 Magicscan" scanner coupled to a" FUJIX Pictography | 3000" printer. Our base computer is always an Apple system upgraded | periodically. | ********** | } In response to Tina's post, I have not seen any | } mention on the list of the scanner I purchased | } a few weeks ago, the Epson Expression 1640XL. | } It has 1600dpi optical resolution (scans at | } a hardware resolution of 1600x3200 dpi) 42 bit | } color (14 bit gray) and Dmax of 3.6. | | I would certainly believe the resolution and the color depth for | this scanner is adequate, but if scanning TEM films is an issue, I'd | seriously advise measuring the optical density of your films ... I've | heard these approach OD} 4 ... which would imply you might consider the | dedicated film scanners, e.g., Polaroid 45 Ultra or the new Nikon | LS-8000. | ********** | I have the scanner you are looking at & like it a lot. To be quite honest | I do not find that I need to exploit it's full capabilities. If I were in | the market again, looking at newer technology I would be interested in a | faster scanner of similar quality. Yes I want my cake & to eat it too :). | I'll give you this analogy. If I have 10 negatives I will franchise my | time, that is let things scan while I hang out in the office doing other | things. If I have 20 negatives, I'll probably go to the darkroom to make | photos. It is quicker & paper is cheaper. BTW I have an Epson 870 inkjet | that produces nice quality images... cost is down to $180 US, (now the | Epson 880)....no financial interest in these companies. | ********** | There was a thread recently on scanners for TEM film. I have looked up | all the models mentioned, on the web and called agents for prices - and | produced a comparative table, given below. | I do not guarantee that the figures are accurate but they are my best | interpretation of the data given. | In the light of experience and Nestor's comments, I would suggest that | 2000 dpi is a minimum for TEM negatives. You may be able to get away | with less nine times out of ten, but there will be occasions when you | need more. | I would exclude the Minolta and all the Epsons from consideration | (despite the incredibly low prices of some of the Epsons) because of the | low pixel density. | Among the rest the Nikon has the best pixel density and the best optical | density (another critical parameter for TEM negatives). The price is | very competitive too. The Nikon web site does not give a time for | scanning a negative. On the face of it the Nikon would be a best buy - | get a separate, inexpensive flatbed scanner for the other work. | These comments are all my own opinions based on manufacturers' data. | Since we are considering purchase any comments to the contrary would be | most welcome. | Code Maker Model Type | A Agfa DuoScan T2500 Flatbed | -Transparency | included | | B Epson 1640 several versions Flatbed | -Transparency | option | 1680 several versions | | C 1600 several versions Flatbed | -Transparency | included | | D Imacon Flextight Precision II Drum -for film and | large | format | | E Minolta Dimage ScanMulti II Film | | F Nikon Super Coolscan 8000ED Film | | G Polaroid 45 Ultra Film | | H Umax Powerlook 3000 Flatbed | -Transparency | included | | | | | | Code dpi OD Time Price | Opinion | at 6 x 9 cm | | | A 2500 x2500 3.4 3 min $4,500 | Fair | B 1600 x 3200 3.6 $300-$3000 | Poor | $800-$1400 | Poor | | C 1600 x 3200 3.3 $650-$1160 | Not suitable | | D 2240 x2240** 3.9/4.1 N/A above $10k | Good: low pixel density | | E 1128 x 1128 3.6 | Not suitable | | F 4000 x 4000 4.2 N/A $2,695 | V. Good | | G 2500 x 2500 3.8 5 min $7,495 | Good but pricey | | H 3048 x 3048 3.6 3 min $6,499 | | ********** | I too am about to buy and I would make a couple of comments on your | evaluation. First, let me remind everyone that the Dynamic range is | a log scale so small numerical differences are significant. | | I also think the Nikon Coolscan 8000 looks great but it only takes a | 2.5 x 3.5 negative which is smaller than my JEOL and Hitachi EM | negative sizes (~ 3 1/2 by 4 1/2"). Have these EM manufacturers gone | to a smaller film size or is Nikon using a non-Japanese EM as their | standard? seems odd but I don't see how the Nikon would be very | useful. You say a {2000 line scanner would be useful 9 out of 10 | times but want the 2000+ lines for the occasional high res scan. I | would argue that the size of the negative was the more important | variable to be worried about. The Nikon couldn't handle 4x5 LM | negatives or transparencies from autoradiography of | Westerns/Northerns, etc. | | My leading candidate is the ArtixScan 1100 has a Dmax of 3.9 (about | $1600 with SCSI card). This was has a 1000 x 2000 dpi resolution. | more details at www.microtek.com. This is my leading candidate. It | was 4 negative carriers and I await word whether one could be | modified to carry a 3 1/2 by 4 1/2 negative. At worst, I will have | my scientific instrumentation shop guys fabricate a holder. It comes | with a glass 8 x 10 glass carrier for odd size negs but I want to | avoid Newton rings and want a glassless carrier. | | I would appreciate comments on the following argument (I think I have | this correctly figured out but am not sure since so many out there | seem to want to have a higher resolution scanner). I have a Fuji | Pictrography 3000 printer with a 400 dpi output that is as good as | any other widely available printer in the academic world. If you | figure the maximum published image size is about 8 inches, that would | mean the maximum image size be 3200 dpi wide. A 1000 dpi scan of my | negative would be 4500 x 3500 dpi. I could crop by about 28% or 10% | depending on the orientation of the negative and still be taking full | advantage of the printer resolution. In reality, most EM publication | prints are smaller than 8" wide so one could crop even more and still | not need more than 1000 dpi. A resolution } 1000 dpi would be | useful for subtle morphometric analysis but a 4000 dpi scan of a 3 x | 4 negative would be 192 MB. That is pretty big for doing morphometry | on! A 1000 dpi scan of a 3.5 x 4.5" negative would be about 16 MB | and that is much more manageable. Perhaps the difference is in the | type of EM we are doing. I am working with biological specimens | doing standard thin section type stuff. are you doing some Material | Sci application that demands more? | I would love to take advantage of the Firewire option but my | information is that the 8700 has a Dmax of 3.4 vs the 3.9 for the | 1100. That is a significant difference. Do EM negatives of | biological thin sections reach that? I think so. I do a lot of EM | immunocytochemistry and have to look for gold (intensely black) | against a very dark tissue component so I am hoping the higher Dmax | improves my results. I frequently scan negatives on a Umax 1100 | (Dmax 3.4??) and can't differentiate the gold from the background | although by eye I can discriminate them when the negative is placed | on a light box. Changing my exposure would give me an unusable | image for the rest of the tissue. Maybe this is an extreme case but | I suspect that lots of "dark organelles" (e.g., lysosomes, nuclei) | have fine structure that get lost in the scanning with a low Dmax | scanner. | ********** | Your information is correct and mine is not. The Dmax of the 8700 is 3.4. | ********** | A colleague and I each recently bought Microtek scanners to scan TEM | negatives. I have the Artixscan 1100 and he has the Model 8700 which has | similar characteristics (actually higher resolution -1200dpi), 3.9 dmax at | 42 bits color (14 grayscale), and the glassless film carrier setup. The | 8700 has USB and Firewire interfaces and is cheaper ( {$1000), and the 1000 | dpi Model 1100 has a SCSI interface. You might want to check out the | specs of the lower cost model 8700 on the microtekusa website if your | computer can handle USB or Firewire. | Both scanners have performed up to our expectations, which I would | characterize as modest. Microtek does not supply a 3-1/4 x 4 " negative | carrier for standard size TEM film but you can easily make a serviceable | one from stiff paper or light cardboard. | | How much scanner resolution should you buy? The answer depends on how you | intend to use it. Most applications do not require capturing the full | resolution of the negative. From a practical viewpoint, the scanner | resolution just determines how many times you can magnify the negative | image to produce the final print size. For example, to get a | publication-size print at 300 dpi, an image scanned at 1200 dpi scan could | be zoomed 4X. A practical alternative to spending more for higher | scanning resolution is to take photos at higher | magnification. One exception is with lattice images from the TEM, which | (depending on the lattice fringe spacing on the negative) might require | higher scan resolutions to avoid getting a moire effect. (Of course, not | everyone agrees. My colleague prefers to always scan at the maximum | resolution). | | What does a Dmax of 3.9 mean to you? To me it means a very dark | negative. D is the log of the transmitted to incident intensity ratio. I | wonder if users ever actually verify the manufacturer's specs with a | calibrated density target. A Dmax of 3.9 can be useful for scanning TEM | diffraction patterns that might have high contrast, but TEM micrograph | negatives of metals and ceramics generally don't have that much contrast | and biological thin section photos tend to have rather weak contrast. If | your negatives are simply dark, use shorter photo exposure | times. Scanning with maximum allowed grayscale resolutions | (e.g., 14 bits rather than 8) is highly recommended if you intend to | enhance or adjust images, but that's another story. | I believe that those Agfa scanners are OEM by Microtek. If budget is the | concern, I would recommend buying a Microtek Scanmaker 5 ($1,100) instead, | which I have used for scanning quite a few EM negatives and have | satisfactory results. The Dmax and the dynamic range for Scanmaker 5 is | are about 3.7 and 3.4. Another model your colleague might consider is an | Agfa Duoscan HiD ($ 2,500) which has a higher Dmax of 3.7, but less | optical resolution of 1,000 dpi compared with 2500 dpi on a DuoScan | T2500. What others failed to mentioned is that the DuoScan T2500 only has | a narrow strip on the CCD bay being capable of scanning at 2,500 dpi, | otherwise the true optical resolution is 1,000 dpi. Although a lot of | investigators think that the higher scanning resolution, the better, my | personal bias is leaning toward to purchase a scanner having at optical | resolution at 1,000-1,200 dpi. Umax also carries a few mid- to high-end scanners such as | Powerlook III for routine negative stains. My personal experience for the | UMAX scanner is only limited to the Powerlook II, a mid-range scanner | which gives more grayish scanned images compared to those high end models | I mentioned previously. However, it is a descent scanner if you are | working with color transparencies. | | ********** | I have not summarized the lengthy thread about film (logarithmic) vs | digital (linear) response! | | | Aloha, | Tina | | ************************************************* | * Tina (Weatherby) Carvalho * tina-at-pbrc.hawaii.edu * | * Biological Electron Microscope Facility * (808) 956-6251 * | * University of Hawaii at Manoa * http://www.pbrc.hawaii.edu/bemf* | **************************************************************************** | | | | | |
Roger Moretz, Ph.D. Dept of Toxicology Boehringer Ingelheim Pharmaceuticals, Inc. 900 Rigdebury Road Ridgefield, CT 06877 203-798-5448
_______________________________________________________ Send a cool gift with your E-Card http://www.bluemountain.com/giftcenter/
Question: I am trying to do immunohistochemistry on lung tissue. What is the best way to fix this tissue? Is there a good reference for protocols? Thank you
A slip of the finger caused me to send an un-topical joke to all. While no has yet criticized (though some have expressed appreciation and/or agreement), I feel that the joke was inappropriate to the venue. My apologies.
Go for it! The "magic" loop is great and makes life sooo much easier. I use it for collecting both the semi-thin sections (instead of an eye lash) and the thins. Not only you get better control of the position of sections on a grid, it also saves you time. Although the loop requires very careful handling - it withstands only certain number of accidents when you bend it from its original angle - it's worth every penny. Good luck, Alice.
-----Original Message----- } From: Claudia Hayward-Costa To: Microscopy-at-sparc5.microscopy.com Sent: 4/25/01 2:57 AM
Dear Microscopists,
I wondered how commonly people use a tool (like a loop) to pick up ultrathin sections.
Does it make life easier or does it only introduce different problems?
I was told that I can buy some "luxury" - but am still undecided.
Your views would be very much appreciated.
Regards
Claudia
Dr. C. Hayward-Costa School of Life Sciences Kingston University Penrhyn Road, Kingston upon Thames Surrey KT1 2EE, UK 44(0)208 547 2000 x 2240 Email: c.hayward-at-kingston.ac.uk Fax: 44(0)208 547 7562
If you are lucky, then you can find the FWBell4080 still with a distributor. However, it was discontiuned. However, you can still get the 4090, which has nice output.
If you also go for the Extech sound meter (as per Nestor's suggestion), then get the high end one. Model 407355 has software for graphing the results. It is well worth the extra $.
JQ
} } From Microscopy-request-at-sparc5.microscopy.com Wed Apr 25 05:22:50 2001 } From: "Anaspec" {anaspec-at-icon.co.za} } To: "'John C. Wheatley'" {John.Wheatley-at-asu.edu} , } {Microscopy-at-sparc5.microscopy.com} } Subject: RE: Search Coil } Date: Wed, 25 Apr 2001 07:46:27 +0200 } } } Hi John } The best bet is to contact your local RS electronics supplier ( find them on } http://www.rs-components.com/ )and ask for the ELF Magnetic Field Strength } meter part number 212 837. } We use it extensively for site tests and have found it to be just as } accurate as using a professional test kit. The only difference between this } unit and a search coil to a scope is that this device simply tells you if } you have a field problem between 20 to 1200Hz where a more expensive search } coil will tell you exactly what frequency it is that is causing the field. } } Good Luck } Luc Harmsen } Anaspec, South Africa } Technical support on microscopy. } Tel + 27 (0) 11 476 3455 } Fax + 27 (0) 11 476 7290 } anaspec-at-icon.co.za } www.anaspec.co.za } } } -----Original Message----- } } From: John C. Wheatley [mailto:John.Wheatley-at-asu.edu] } } Sent: 25 April 2001 01:10 } } To: Microscopy-at-sparc5.microscopy.com } } Subject: Search Coil } } } } } } I need to find a commercial source for a calibrated search coil to check } } stray fields in microscope rooms. Does anyone have any experience with } } purchasing this item? } } } } John C. Wheatley } } Lab Manager } } Arizona State University } } Center for Solid State Science } } PSA-213 } } BOX 871704 } } Tempe, AZ 85287-1704 } } } } } } Phone: (480) 965-3831 } } FAX: (480) 965-9004 } } John.Wheatley-at-ASU.Edu } } } } } } } }
Until recently, I was very much against using a loop to pick up ultrathin sections. A student in the lab started using it and I kind of reluctantly gave it a try. Now I am a regular user of the loop to pick up sections on coated grids.
First I clean the loop with 100% ethanol, air dry and pick up sections. I place a coated grid on a piece of filter paper and bring the loop with the sections down onto the coated grid and let the filter paper soak the water droplet. The sandwiched grids are allowed to air dry for few minutes and you can then remove the loop using a fine tweezer. It works just great.
I bought my loop from Electron Microscopy Sciences. No financial interest with EMS.
Just give it a try.
Soumitra
************************************************************* Soumitra Ghoshroy Ph.D. Electron Microscopy Lab and Fluorescence Imaging Facility Graduate Faculty, Department of Biology Box 3EML New Mexico State University Las Cruces, NM 88003 Tel: 505-646-3600 Fax: 505-646-5665 e-mail:sghoshro-at-nmsu.edu http://confocal.nmsu.edu/eml
On Wed, 25 Apr 2001, Claudia Hayward-Costa 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 Microscopists, } } I wondered how commonly people use a tool (like a loop) to pick up } ultrathin sections. } } Does it make life easier or does it only introduce different problems? } } I was told that I can buy some "luxury" - but am still undecided. } } Your views would be very much appreciated. } } Regards } } Claudia } } Dr. C. Hayward-Costa } School of Life Sciences } Kingston University } Penrhyn Road, Kingston upon Thames } Surrey KT1 2EE, UK } 44(0)208 547 2000 x 2240 } Email: c.hayward-at-kingston.ac.uk } Fax: 44(0)208 547 7562 } }
In a message dated 4/24/2001 9:41:00 PM Mountain Daylight Time, John.Wheatley-at-asu.edu writes:
} I need to find a commercial source for a calibrated search coil to check } stray fields in microscope rooms. Does anyone have any experience with } purchasing this item?
An inexpensive (~$90) digital gauss meter (Extech Model #480823) that is quite adequate for locating sources of 30 to 300 Hz magnetic fields can be found at Meters and Instruments, "www.MetersandInstruments.com", (800) 773-0370.
Joe _________________________________________ Joe Nabity, Ph.D. JC Nabity Lithography Systems E-Beam Lithography using Commercial SEMs & STEMs PO Box 5354, Bozeman, MT 59717 USA Voice: (406) 587-0848 FAX: (406) 586-9514 E-mail: info-at-jcnabity.com Web: www.jcnabity.com
Picking up from above is easy with the grid held by a forceps (I prefer a curved Dumont 5 or 7 with a rubber "clamp" to hold it together), the sections just "jump" on my filmed grid. I have tried a Perfect Loop (commercially available) and I also liked it - I could see no difference in "section quality" comparing both methods. Picking up with a grid from below the water surface I do not like (especially with filmed grids, but thats just my personal opinion)
I sometimes loose sections when I am not carefull enough with my eyelash but very very rarely when picking up...
Of course for cryosectioning a loop is an absolute must - and here I clearly prefer the Perfect Loop! The droplet of sucrose solution it can hold is much larger than in a homemade wire loop (gives you a bit more time in the cryochamber) and you can nicely use this large drop as a magnification lens for locating your cryoscetions before picking them up.
Best regards,
Joachim
Dr. Joachim Prutsch Product Manager EM Specimen Preparation
This reminds me - we used to get Epon or Spurr's off sandwiched glass slides by putting them in -20C freezer for 1/2 hr and then work the sample off. John Shields EM Lab Univ. of GA Athens
On 24 Apr 2001, at 14:47, Douglas Keene 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 } ---------------------------------------------------------------------- } -. } } } } You may want to try dipping just the thermanox portion of } your block into liquid nitrogen. The sudden change of } temperature will likely loosen the thermanox away from your } sample. I do not expect epon to be a problem, but I do } know that it works well with Spurrs. } } Good luck, } } Doug } } On Mon, 23 Apr 2001 10:12:10 -0500 } "tbargar-at-unmc.edu"-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, } } I need advice on embedding Thermanox coverslips. It's } } supposed to peel off leaving the monolayer behind, but I'm } } not having much luck. I'm using Aralidite 502 as the } } embedding medium. Would a another resin work better? I } } would appreciate any and all advice. Thanks. } } } } Tom Bargar } } EM Lab } } UNMC } } 402-559-7347 } } tbargar-at-unmc.edu } } } } } } ---------------------- } Douglas R. Keene } Associate Investigator } Shriners Hospital Research Facilities } 3101 S.W. Sam Jackson Park Road } Portland, Oregon 97201 } phone: 503-221-3434 } FAX: 503-412-6894 (9-5 PST) } e-mail: DRK-at-shcc.org } }
Hi Listers, What's the best 'glue' to use for semiconductor cross section sample prep? We use Gatan G1 to stick the 'sandwich' together (bake at about 150C for an hour) BUT then we stick a 3mm supporting washer onto the mechanically thinned sample using Devcon 5 Minute epoxy prior to ion beam milling. In our constant search for less 'contamination', we are not sure if this is good practice in a FEG (especially) TEM. Are there any other alternative 'glues' that would do the job - without baking? Alan Walker
********************************************* Alan Walker Dept of Electronic and Electrical Engineering University of Sheffield Mappin Street Sheffield S1 3JD United Kingdom
A quick question for the clinical diagnostic people out there. I was wondering what an average turn around time would be for an EM specimen. that is from time of arrival to the time it is placed in the scope.
{HTML} {FONT FACE=arial,helvetica} {FONT SIZE=2} A quick question for the clinical diagnostic people out there. I was {BR} wondering what an average turn around time would be for an EM specimen. that {BR} is from time of arrival to the time it is placed in the scope. {/FONT} {/HTML}
I am trying to find a place to donate my father's microscope. He was a physician, born in Germany. The microscope was bought in Germany, probably in the 1920's or 1930's. It is stored in a wooden case. Is there any place this instrument could be useful? A school or university in the third world? A school in the US?
Please contact me, offline, at ulmithaca-at-home.com.
The Microscopy and Flow Cytometry Core Facility within the MRC Clinical Sciences Centre, Imperial College School of Medicine, has immediate openings for:
Microscopy Research Associate/Assistant (Ref: MRA) Microscopy Core Facility
The imaging of molecules within cells, organs and whole organisms is at the forefront of scientific developments in the life sciences. The Clinical Sciences Centre (CSC) is a leading research institute with groups developing new avenues in the areas of confocal laser scanning microscopy (CLSM), live-cell imaging, scanning ion conductance microscopy, position emission tomography (PET), and nuclear magnetic resonance (NMR). The MRC is seeking a highly motivated individual to be responsible for this state-of-the-art microscope core facility at the CSC, to provide full support for confocal, live-cell imaging, and other fluorescence microscope users within the institute. The successful candidate should have a science degree in a relevant subject. Experience in confocal, live-cell and digital imaging (particularly Leica confocal microscope and/or deconvolution software) would be highly desirable but not essential, as training can be given. Duties will include direct technical support, training and supervision of users. Involvement in research project using fluorescence microscopy will be encouraged (further information about research at the CSC can be found at www.csc.mrc.ac.uk). Informal enquiries and further information about the position to Drs Ana Pombo (tel: +0/44 20 8383 8232; ana.pombo-at-csc.mrc.ac.uk) or Alex Sardini (tel: +0/44 20 8383 8270, a.sardini-at-csc.mrc.ac.uk), MRC Clinical Sciences Centre, Imperial College School of Medicine, Hammersmith Campus, Du Cane Road, London W12 0NN. The closing date for applications is 30 April 2001.
Research Assistant (Ref: FCRA) Flow Cytometry Core Facility
Flow cytometric analysis and cell sorting are important for many areas of research in the CSC, including immunology, gene expression and stem cell work. The facility is operated jointly with Imperial College School of Medicine (ICSM) and consists of 2 FACS Vantage cell sorters and several bench top analysers. The MRC is seeking a full time research assistant to help our existing staff provide a cell sorting and analysis service , advise users on the design of their experiments and liase with service engineers (all instruments are on full service contracts). Involvement in research projects will be encouraged. The successful candidate will have a strong interest in the interface between biology and technology. Experience in flow cytometry will be an advantage but is not essential as training can be given. Informal enquiries and further information about the position to Dr Matthias Merkenschlager (tel: +0/44 20 8383 8236/9; matthias.merkenschlager-at-csc.mrc.ac.uk), MRC Clinical Sciences Centre, Imperial College School of Medicine, Hammersmith Campus, Du Cane Road, London W12 0NN. The closing date for applications is 30 April 2001.
The CSC is an institute funded by the Medical Research Council and forms part of the ICMS, based at the Hammersmith Hospital in West London. This recently established centre has first class facilities and provides investigators from clinical and basic science backgrounds with the opportunity to pursue innovative, multidisciplinary research within the established clinical base of ICMS. Salaries will be on the MRC's own pay scales and will be commensurate with experience. Further details on how to apply are available from the Human Resources Group tel: 020 8383 3446/7, quoting the relevant reference above.
Advertised in Nature and New Scientist.
Ana Pombo, D.Phil. Nuclear Organisation Group MRC Clinical Sciences Centre Imperial College School of Medicine Hammersmith Hospital Campus Du Cane Road London W12 0NN UK
Although the 1.2MeV HVEM at ANL has sadly departed, we at NCEM want to remind everyone on the listserver that there still exists a HVEM in the US available for on-site and remote use, free of charge with approved proposal.
The Kratos 1.5 MeV HVEM has recently been refurbished (following some substantial high voltage and vacuum issues). It is presently operational at 1 MeV, and is being conditioned for use at 1.5 MeV. 1.5 MeV operation is expected by the end of June, at the latest.
NCEM is actively seeking user proposals for this instrument. Please see:
http://ncem.lbl.gov
http://ncem.lbl.gov/frames/hvem.htm
for further details regarding the instrument's many capabilities (among them straining stage experimentation and environmental cell work), or feel free to contact either myself or Doug Owen - DKOwne-at-LBL.gov
Regards, Eric Stach -- Eric A. Stach Staff Scientist National Center for Electron Microscopy Mail Stop 72-150 Lawrence Berkeley National Laboratory Berkeley, CA 94720 Phone: 510.486.4634 Fax: 510.486.5888 http://ncem.lbl.gov
I am trying to find a place to donate my father's microscope. He was a physician, born in Germany. The microscope was bought in Germany, probably in the 1920's or 1930's. It is stored in a wooden case. Is there any place this instrument could be useful? A school or university in the third world? A school in the US?
Please contact me, offline, at ulmithaca-at-home.com.
Dear Alan, In the original semi-conductor cross-section discussions, M-Bond 610 was recommended. It is a srain-gauge glue, two-part, that requires about 100 deg. C to harden, but it is low viscosity and left a very thin joint. When we were waitng for our order to arrive, we used Devcon 2-ton epoxy, which is higher viscosity and requires eight hours to harden, but still made a thin joint if you squeezed it in a parallel-jawed vice. Once these are hard, we saw no evidence of out-gassing, but we do not have a FEGTEM. At 04:55 PM 4/25/01 +0100, you wrote: } } Hi Listers, } What's the best 'glue' to use for semiconductor cross section sample prep? } We use Gatan G1 to stick the 'sandwich' together (bake at about 150C for an } hour) BUT then we stick a 3mm supporting washer onto the mechanically } thinned sample using Devcon 5 Minute epoxy prior to ion beam milling. } In our constant search for less 'contamination', we are not sure if this is good } practice in a FEG (especially) TEM. Are there any other alternative 'glues' } that would do the job - without baking? } Alan Walker
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
Technically you can not remove this type of plastic from a section. Several methods have been published and suggested however, they often destroy the section at the same time. Pam Marcum
-----Original Message----- } From: Rinaldo Pires dos Santos [mailto:rinaldo-at-ufrgs.br] Sent: Wednesday, April 25, 2001 8:23 AM To: Listserv Microscopy
Hi all,
Is there some procedure for to remove hydroxyethylmetacrylate resins (Leica Historesin) from semithin sections (1-2 micrometers)? Thank you.
Dr. Rinaldo Pires dos Santos Lab. of Plant Anatomy - Dept. of Botany E-mail: rinaldop-at-uol.com.br UFRGS - Porto Alegre - RS Brazil
If you are not doing site-specific work with semiconductors as I assume that you are from your message, I would suggest that you use the small angle cleavage technique to do your cross sections. I have done III-V and Si based samples with superb results. Take a look at John McCaffrey and my article in the Number IV MRS TEM Sample Prep book, Vol 480, 1997 for a detailed procedure. South Bay Technology sells the "Micro Cleave Kit". This technique is inexpensive and gives great samples. I have used the samples with no plasma cleaning in a FEGTEM without any problems. In addition, you have no ion milling damage. In fact for a FEG, you have a disadvantage because there is little or no amorphous area to use for focusing.
I use the H-22 silver epoxy from Epoxy Technology to put the samples on to the grids. This requires heating. When I do not want to heat the samples, I have used a Duro 90 minute epoxy for long working times. Unfortunately, it takes about 12 hours to fully cure. I have also used these samples in a FEGTEM without problems. I have cured them in atmosphere and in a nitrogen atmosphere.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center P. O. Box 11472 (letters) Guys Run Rd. (packages) Pittsburgh, PA 15238-0472
Walck-at-PPG.com
(412) 820-8651 (office) (412) 820-8515 (fax)
-----Original Message----- } From: A.Walker [mailto:Alan.Walker-at-sheffield.ac.uk] Sent: Wednesday, April 25, 2001 11:56 AM To: Microscopy-at-sparc5.microscopy.com
Hi Listers, What's the best 'glue' to use for semiconductor cross section sample prep? We use Gatan G1 to stick the 'sandwich' together (bake at about 150C for an hour) BUT then we stick a 3mm supporting washer onto the mechanically thinned sample using Devcon 5 Minute epoxy prior to ion beam milling. In our constant search for less 'contamination', we are not sure if this is good practice in a FEG (especially) TEM. Are there any other alternative 'glues' that would do the job - without baking? Alan Walker
********************************************* Alan Walker Dept of Electronic and Electrical Engineering University of Sheffield Mappin Street Sheffield S1 3JD United Kingdom
The loop works great!!! It's kind of pricy, so careful handling is a must, but it can be a real headache saver. I, too, bought my loop from Electron Microscopy Sciences. I have no financial interest in EMS.
Good luck,
Elizabeth P. Bray Plant Chemist, Central Laboratory South Carolina Electric and Gas Co. 2102 N. Lake Dr. Columbia, SC 29212
----- Original Message ----- } From: "Claudia Hayward-Costa" {LS_S562-at-crystal.kingston.ac.uk} To: {Microscopy-at-sparc5.microscopy.com} Sent: Wednesday, April 25, 2001 5:57 AM
I rather enjoyed it.
Earl
----- Original Message ----- } From: "Chuck Butterick" {cbutte-at-ameripol.com} To: {Microscopy-at-sparc5.microscopy.com} Sent: Wednesday, April 25, 2001 5:36 AM
SUMMER I 2001 COURSE ANNOUNCEMENT - Transmission Electron Microscopy (BIO. 221-Section B)
NASSAU COMMUNITY COLLEGE, Garden City, Long Island, New York
A five week, Summer Session I 2001 semester, course in Biological Transmission Electron Microscopy is being offered by the Biology Department of Nassau Community College. This is a 4 credit course offered four days per week (Monday through Thursday) between the hours of 8:00 am and NOON. Classes will begin on May 29 and end on June 28, 2001.
This is a "hands-on" course emphasizing biological specimen preparation, ultra-thin sectioning involving block trimming, glass knifemaking and operation of the ultramicrotomes (Sorvall MT-2B and LKB Ultrotome III), thick and ultra-thin section mounting and contrast staining (UA and Pb citrate), grid support films (formvar, carbon), student operation of the TEM (Hitachi HS-8, Philips EM 300) and production of electron micrographs through the process of black & white photography, and electron micrograph analysis. Students will work on a chosen sample(s) with the goal of producing a portfolio of high quality TEM photomicrographs of that sample(s).
The course is widely transferrable and the cost per credit is reasonable at $92 per credit (for Nassau County residents or New York State residents with a certificate of residency).
More information about the Bio-Imaging Center at NCC, course descriptions and syllabi, and the beginnings of a student gallery of EM photomicrographs is available at our web site. The URL is {http://www.sunynassau.edu/webpages/biology/becks.htm} .
Interested individuals should register as soon as possible since the course is limited to a total enrollment of ten (10) students.
If you have further questions, you should e-mail me directly at the address below.
For information about mail or telephone registration (Dial-a-Course) point your browser to http://www.sunynassau.edu/courses/sum01/away.pdf. The phone registration option is available until tomorrow 4/26/01 (6:30 PM) by calling 516-572-7131 or 7372 or 7425.
P.S. A Fall 2001 TEM course is also being offered (BIO 221 - Section E2) on Thursday evenings beginning at 5:30 PM.
Stephen J. Beck Associate Professor Bio-Imaging Center/Electron Microscopy Department of Biology Nassau Community College Garden City, NY 11530 Voice Mail: (516) 572-7829 Email: {becks-at-sunynassau.edu} URL: {http://www.sunynassau.edu/webpages/biology/becks.htm}
Dr. Rinaldo Pires dos Santos wrote: ================================================================= Is there some procedure for to remove hydroxyethylmetacrylate resins (Leica Historesin) from semithin sections (1-2 micrometers)? Thank you. ================================================================== Perhaps there are other methods, but one that has been used quite successfully has been the use of plasma etching in a unit such as the SPI Plasma Prep™ II.
Some examples are on the SPI Supplies website. For example, on URL http://www.2spi.com/catalog/instruments/etchers1.html you will see an example of a thick section: "Statocyst organ etched 3 min. using O2", and of a thin section, "Low temperature oxygen plasma etched thin section of bacterium embedded in SPI Chem Low Acid GMA for TEM."
Like with any technique in LM or EM, this approach has its limitations. But it is a good way, at least in some instances to remove GMA without disturbing the rest of the sample.
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI SPI SUPPLIES FAX: 1-(610)-436-5755 PO BOX 656 e-mail: cgarber-at-2spi.com West Chester, PA 19381-0656 USA Cust. Service: spi2spi-at-2spi.com
Look for us! ############################ WWW: http://www.2spi.com ############################ ==================================================
Thanks to all that responded to my question about making it easier for students to cut thin sections using glass knives. Three key points emerged: small block faces, fresh knives (changed often), and a variety of different embedding media. A summary of the responses is below.
Thank you again.
Dick Briggs 1. This is I think a quite common problem. The students in my lab normally embed in Spurr's firm (both animal and plant tissue) and they initially use glass knife and after they feel comfortable with glass knife, I let them use a diamond knife to do their final sectioning. We also use MT2 and MT2B ultramicrotomes.
2. I think LR White resin should fit the bill. It penetrates easily and you don't have the same level of toxicity as with Spurr's (which is always good for beginners).
3. Teaching microtomy is the most difficult portion of any microscopy class. There is such a long learning curve and so many pitfalls that getting good sections depends on the ultimate patience and dexterity of the student. Some will get it right off, others must persevere. You probably already know my suggestions but I'll put my 2 cents in. My suggestion is to start the student with a quality block of a material that cuts easily. Mouse or rat, kidney or liver are good candidates. As they get good sections on these then move them to tissues that may be more difficult. They may be able to troubleshoot problems without questioning the block. I would also ask how the knives are broken? It may be a dull knife problem. A balanced break really helped improve the quality of the knives we were getting from an LKB knife breaker. Additionally a slow break has always seemed better. Wetting the scribe before breaking also lessens the breaking force required and improves edges. Of course small block faces are also going to improve the results dramatically but trying to get the student to cut down the block face is difficult. There are all the usual excuses.
4. I think some of the problem is your choice of tissue rather than plastic. Plants have tough walls, and open spaces mixed together. This causes a lot of different plasticities (?) within a section because some areas will infiltrate to a better or worse degree than a neighboring area. You would have much better luck using a homogeneous mammalian tissue like liver.
5. I faced the same problem. I placed most of the blame on the novices' inability to make good knives. My solution: I break about 25-50 knives a week and give them to the students. I have found that it greatly enhances their ability to get good sections. I also got a grant to buy a new ultramicrotome (Ultracut T). Between my knives and the new 'tome, things are much better now.
6. I think that glass knives dull very rapidly (either damage or plastic build-up on the edge), and so we have always tried to get the first few sections at a given place on the knife edge, which tend to be the best. I wrote a brief procedure for students years ago, based on that approach, and have typed it below (the handout had a drawing of a glass knife, with area A = 1/3 of edge on the side where the whorl meets the edge, area B = middle 1/3 of knife edge, area C = 1/3 of edge at side where whorl is farthest from the edge):
a. Face the block in area C of knife (see diagram). Cut semithin sections (if desired) in area B of knife. Then move to area A. b. Bring the block face parallel with the knife, using the shadow method. Then use the shadow to bring the block face as close to the knife (but without touching) as you dare. c. With the ultramicrotome set for ultrathin sections, manually turn the microtome wheel quite rapidly until you see the first sign of contact (usually a sliver off one side of the block face), then stop turning. d. Turn on automatic sectioning at usual slow cutting speed. Cut about6-12 full-face sections, then stop and pick them up on EM grids. e. Retract the stage slightly, move laterally to another place in area A, and repeat steps 2-4. Continue until area A has all been utilized (or until you have all the sections you need).
7. I teach EM at a four-year college also. I use Spurr's and glass knives made with an LBK 7800 knife maker. I am careful not to select difficult tissues (nervous or muscle or bone or cartilage), but generally my students have been able to get excellent sections by the end of the semester. a. Details to consider: Do your knife maker and microtome work properly (I have an MT2-B and a newer Leica UltraCut. Some students actually prefer the MT2-B. Are you fully dehydrating specimens and are you using truly dry acetone to dehydrate--I dehydrate mine with ashed CUS04. b. Are you buying good float glass and are the students cleaning it fully and handle it carefully. Mine wash with detergent and hot water, DI water, and ethanol, and wash glass and make knives immediately before use.
8. I always have good results using Spurr's. The sample morphology is not as good as other resins. Be sure to polymerize for a good 48 hours or more.
9. I'm doing the same thing with my EM class this semester. I use only Spurr's for both animal and plant tissues and glass knives (our knife breaker is as old as our MT-2). I find students have greater success with the smallest block faces. I also have them use Formvar-coated grids (100 mesh) that tend to stabilize those sections with bad knife marks. I start by making them get thick sections on larger block faces -- they learn the mechanics of the microtomes, block trimming, etc. When they are ready to get thin sections it seems easy to them to just let the motor take over.
10. I typically use an Epon-Araldite mixture for all of my biological samples (generally mammalian soft tissue samples) without any sectioning problems over many years. For botanicals you might need a low viscosity resin like Spurr's. I use MT-2B's in teaching of my students and they rarely have problems obtaining good sections (silver) in ribbons. The key I believe is in the block trimming. My students trim their trapezoid faces no larger than 0.5mm on the longest side (many are around 0.25 mm). I also have the students use the flimsy double-edge razor blades vs. the single edge variety - a trick I learned from my mentor. The blades are much sharper (and more difficult to handle - I require my students to have Band-Aids on hand for block trimming ;-) and give much smoother top and side of the pyramid surfaces. The Epon-Araldite Mixture I use is as follows:
Stock Solution (can be frozen) Small Volume Large Volume Araldite 6005 12.5ml 25ml Poly/Bed 812 15.5ml 31ml Dibutyl phthalate 2ml 4ml
Final Working Solution
Stock Solution 4ml 8ml DDSA 10ml 20ml DMP-30 14 drops* 28 drops* (*Drops introduced with a Pasteur Pipette) 11. Centuries ago, I used the following mixture to embed cellular slime molds for thin-sectioning with glass knives: Araldite 502 13.5 ml DDSA 11.5 ml DMP-30 0.4 ml The Araldite made it a little softer and easier to cut. Hope it helps!
12. My recommendation is to use LR White on a tissue like spleen, kidney, intestine or heart muscle. One mouse would give you more than enough tissue to last a lifetime. Plants are notoriously difficult to section (walls, air spaces, etc.). If you wish to avoid killing animals, you might sacrifice one mouse and put away tissues in fixative or buffer and then provide the students with vials containing the tissues. Also, sometimes vivariums may be euthanizing animals or a fish may die, etc. providing some valuable tissues.
13. When I was an undergraduate (eons ago) I used the "American Araldite" recipe:
Araldite 502 27 parts DDSA 23 parts DMP-30 1.5-2.0% (accelerator) I never included dibutylpthalate.
We mixed it in old (but clean) baby food jars by volume. We had a "reference" jar with marks at the appropriate places and just lined up the reference jar with the one we were measuring into. Very crude but we got good results. I used small pieces of tissue (kidney, spleen, sm. intestine) and went straight from 1:1 to pure resin and into the oven. The mixture is very viscous but can be warmed a bit to make handling easier. I don't know if plant material would work. I used glass knives and an MT-1, but not after a lot of coffee or a night on the town.
14. I have played this game for 32 years now and still don't consider myself an expert because things fool me all the time. I believe almost any embedding medium will work, if instructions are followed and components are fresh. I use pre-mix kits from TAAB, add bottle one to bottle two etc. It's for an easy life. The leftovers, which can be most of the bottle, go into the freezer. Fresh Spurr's cuts like a treat, sometimes thawed Spurr's is fine, sometimes it is not. The reasons can be maybe a little humidity gets in if not up to room temperature (maybe) or it begins to age (increase viscosity due to starting to cure). Anther things to remember are the smaller the block, the better it cut? Also, don't dwell on one piece of edge for too long, often I get just a couple of grids and then move along. I used to make a special mix which was magic (really). It was TAAB-Hard. With about 1% extra MNA (used to weigh all the components) and 1% DMP-30 accelerator. Curing was also important - 48 hours at 35 degrees C, 24 at 45 and 24 at 60 degrees. It was very hard and brittle. It was sterically inhibited i.e. not highly cross-linked but with as much hardener (MNA) as ever allowed. It cut all day on one knife and was terrific. But people didn't want to follow the curing schedule - 4 days!! But the results were the best sections I ever used to cut; I don't do so well these days, even with diamonds.
15. I too teach an undergrad EM course (with Sorvall MT-2s). I've discovered my students are very reluctant to change knives once started. Forcing constant changes has improved the sectioning. We use Spurr's: 10 ERL 5 DER 25 NSA 0.5 DMAE These proportions work best. After dehydration through 50/75/80/95 we go through 4 100% rinses then 1resion:2 ETOH for 3 hours, then 1ETOH to 2 resins (2-3 hours) then full resin overnight - then bake. I assume you're using these proportions so if you want more detail, let me know. The strange thing is my kids do quite well at getting thins with glass knives - they consistently surpass my expectations.
16. We us Epon with mouse liver. Spurr's would probably be better with plants. Plants are always are a tough go for the uninitiated (though probably easier to obtain and no animal right's issues). Plants also require longer infiltration times and smaller increases between infiltration steps (I usually go 25, 50, 70, 80, 90, and a couple of 100% before polymerization). LR White has been suggested, but as the resin is more hydrophilic it is actually more difficult to learn on at first. The block face has a tendency to wet easily and usually students get frustrated trying to figure out why the block face is getting wet even with the more hydrophobic resins like Spurr's or Epon.
17. I find the same problem. I also find that having them get very close to really mastering thick sectioning first (thereby seeing some really pretty LM slides) greatly helps their confidence, and I also have them use the tiniest faces they can manage (0.1mm?) to get started with thins. They must be the correct shape with no ragged edges (such as you might get after taking (or trying to take) many thicks. Also make sure their knives are good - especially FRESH - and that they are using the correct edge and don't' try to take too many sections before switching knives. Have them start with something like liver where the tissue/resin interface is not an issue. I have tried Spurr's but it is so nasty to use, and is not as stable or as pretty in the EM.
Secondarily, I keep troubleshooting sheets taped to the wall - check size! Check shape! Check knife-edge! Check speed! Check angle! etc. I find this helps beginners understand that lots of factors need to be satisfied - it seems that so many of them probably get by in other courses by cramming the night before, which just doesn't' work here... although some of the 'rules' (such as face size) can be relaxed once they get the basics down.
18. There is a resin additive sold by Electron Microscopy Sciences, called "Sure-Cut Surfactant" (their #21630). I have not tried it myself, but remember a posting on this server 1-2 years ago praising this additive very highly. The guy was saying it saved his students a lot of frustration. There is a trade-off, of course, -- they say this additive somewhat interferes with staining. I would give EMS a call; they are usually helpful. (I have no interest in their sales, of course.)
19. Is it the embeddment or sectioning which is the problem? For sectioning try to keep the section area as small as possible. If you keep the sectioning width less than ~.25mm it makes sectioning much easier.
20. There is no magic embedding media that allows easy sectioning with glass knives. All they can do is trim the blocks as small as possible, and I do mean small. If the block face looks big under the high-end magnification of the stereomicroscope it is still too large. Have them practice (a lot) trimming blocks. Don't them rush into thick sections too soon. Make sure they can master the cutting one-micron sections and them trimming to the desired area. The guy I trained under had no clue. He had us cutting as soon as possible. The drop rate for the class was around 50%. And console them even the big folks had to learn using glass knives. It is possible to do. If they think that is hard wait till they stain the sections with Reynolds lead for the first few hundred times. Tell them to cheer up; at least they are not learning on an MT1 manual microtome like I did.
21. Is your course specifically in electron microscopy or would slightly thicker sectioning be almost as good from the microtechnical perspective? For light microscopy, I embed plant tissue in Glycol Methacrylate (GMA) from Electron Microscopy Sciences. The embedding protocol is simpler, and there is the potential that the plastic is less carcinogenic than Spurr's resin. I have recently completed a project in which I was sectioning largish block faces of plant apices embedded in GMA using a glass knife on an Olympus rotary microtome. Sections of less than 1 micrometer were no sweat and I bet they'd be even easier on an ultramicrotome. Very satisfying from the teaching side as it should keep the student frustration level down. GMA is also quite easy to stain with e.g. Richardson's stain or methylene blue. Unfortunately, GMA is a bit too soft to section thinner and it doesn't hold up well under the e-beam so for electron microscopy I use Spurr's or LR White.
22. I noticed your comments on the Microscopy ListServer about students and thin sections. There are a couple of fairly easy ways to make it less painful. My old graduate advisor and I have worked together for many years and I have watched, and at times helped him teach his EM course. For his students sectioning was always the worst experience. Two things might help: 1) use a soft to very soft formulation of one of the Epon substitutes. In that way trimming and thick sectioning are easier and glass knives do not dull as quickly nor do they need to be perfect. The second is make a small block face. Unfortunately that may be more difficult to do than sectioning. Some friends of mine at UC Davis use the soft formulation of Epon which really makes sectioning and trimming easier. It is soft enough to easily make an imprint with a fingernail.
23. I was a former student at San Francisco State University (I've graduated in May 2000), and I took an electron microscopy course at that university. Back to my electron microscopy course. During part of that course, I was supposed to section some embedded tissue of my choice (heart / liver /? (can't recall off-hand)), and I had considerable trouble using a microtome well. As I recall, we had some Sorvalls available for use, but I'm unsure if they were MT-2s. I prepared all my tissues (and practice, plastic "pellets") by first cutting with straight razor blades to make an acceptable-sized pyramid, then continue by sectioning with a microtome. Glass knives were used on these embedded tissues -- the plastic used was Epon -- and I recall that one major problem I had was in making sure that the microtome sectioned at a consistent speed. In addition, the glass knife edges would dull pretty quickly after a small number of sections were made, and so it became imperative to use several glass knives for any particular embedded material. I eventually was able to section pretty well, but I also picked up sections from below with a grid (of 200 mesh) that I bent rather badly during the collection process. Epon worked OK for sectioning, and while it's true that students can indeed get discouraged, I also know from experience that students have very different aptitudes towards handling microtomes, preparing the pyramids, etc. I hope that my experience with sectioning may help you understand better (from a student's perspective) the various trouble areas that could occur when students are first learning how to section embedded material.
********************* 1st Announcement and call for contributions *****************************
A meeting organised by the Royal Microscopical Society and supported by EMAG, FEI UK Ltd, JEOL (UK) Ltd.,LEO EM Inc., Gatan UK, TVIPS GmbH
DEVELOPMENTS IN ENERGY-FILTERED ELECTRON MICROSCOPY
Wednesday 4 July 2001
Department of Materials, Oxford University
The use of electron energy filters in analytical electron microscopy (EFTEM) is a relatively recent development that is proving to be an extremely powerful tool. In-column filters (e.g. so-called "omega-filters") and post-column filters (e.g. the "GIF") represent different approaches which have their own advantages as well as experimental difficulties. This one day meeting is designed to explore the current state of the art, both with regard to the instrumentation and also applications of EFTEM in life and physical sciences.
Invited speakers: Dr Bernd Feja (Tietz Video & Image Processing Systems GbmH) Energy-filtered electron tomography Prof Joachim Mayer (Aachen University of Technology) EFTEM - the state of the art and future trends Dr Paul Midgley (University of Cambridge) EFTEM image series - taking elemental mapping into a new dimension Prof Michael Trendelenburg (German Cancer Research Center, DKFZ) EFTEM in biomedicine & biotechnology: Recent advances in specific element mapping
* CONTRIBUTIONS Contributed presentations are now being sought. Abstracts of no more than 200 words should be sent by email to: crispin.hetherington-at-materials.ox.ac.uk and should arrive by 31 May, 2001.
* REGISTRATION Registration will be Ł30 RMS/EMAG members, Ł15 students and Ł40 others and a printable form will be avialable from the new RMS website, from May on this page :
http://www.rms.org.uk/current%20events.html#eftem
Further details are available from the organisers: Dr Crispin Hetherington, tel. 01865 273799, crispin.hetherington-at-materials.ox.ac.uk Dr Laurence Tetley, tel. 0141 330 4431, l.tetley-at-bio.gla.ac.uk and from the meeting website : http://www-em.materials.ox.ac.uk/events/eftem.html
Note this meeting is timed to follow the 1-day FEGTEM III meeting to be held on 3 July 2001, also in Oxford (http://www-em.materials.ox.ac.uk/events/fegtem.html) **************************************************************************** ****************
Dr Laurence Tetley Division of Infection & Immunity, IBLS, Integrated Microscopy Facility Joseph Black Building University of Glasgow Glasgow G12 8QQ
OK this is going to seem like a stupid question. I am going to try araldite 502 embedding media. the instruction sheet calls for a volumetric measurement. I would prefer to measure the each chemical in a balance. OK here is the question: do i multiply the specific gravity by the volume or divide it. it has been a long time for me since chemistry 101. thanks
{HTML} {FONT FACE=arial,helvetica} {FONT SIZE=2} OK this is going to seem like a stupid question. I am going to try araldite {BR} 502 embedding media. the instruction sheet calls for a volumetric {BR} measurement. I would prefer to measure the each chemical in a balance. OK {BR} here is the question: do i multiply the specific gravity by the volume or {BR} divide it. it has been a long time for me since chemistry 101. {BR} thanks {/FONT} {/HTML}
If I'm not swamped, which is never, I do a two day processing. Using a Lynx overnight to get into resin I then put the specimens into molds and let them cure for 24 hours. When I come in on day three I can cut thick sections, have a conference with the Doc who submitted the case then cut and stain the thins and I'm ready to sit down at the scope. In reality, A specimen coming into our cutting room is submitted for routine light microscopy and possibility special stains. Those sections are then reviewed and the decision is made to submit the case for EM examination. That can take one or two days. So there are four, possibility five days before I can photograph the case. Another issue is prioritization. I do live patients first and I do the tumors before genetics cases. It makes no sense to work on a possible mitochondria disorder when there is a kid with a chest wall tumor or a brain tumor. The same goes for a kidney biopsy over a ciliary disorder. I have to prioritize and frequently cases get put off a few days until the heats down. With a current volume of about four hundred cases a year and an additional two hundred cases for research I'm often asked "How long will this take?" My usual answer which includes the scoping and printing or making a CD is five working days. Autopsy's are not high on my list and periodically I'll hear that the resident who submitted the case has rotated into another department before the EM is done. To answer your question I'd have to know what your case load is and how many hands are available.
Howard Mulhern Supv. Path. EM Facility Children's Hospital Dept. of Pathology 300 Longwood Ave. Boston, 02115 Ma.
********************* 1st Announcement and call for contributions *****************************
A meeting organised by the Royal Microscopical Society and supported by EMAG, FEI UK Ltd, JEOL (UK) Ltd.,LEO EM Inc., Gatan UK, TVIPS GmbH
DEVELOPMENTS IN ENERGY-FILTERED ELECTRON MICROSCOPY
Wednesday 4 July 2001
Department of Materials, Oxford University
The use of electron energy filters in analytical electron microscopy (EFTEM) is a relatively recent development that is proving to be an extremely powerful tool. In-column filters (e.g. so-called "omega-filters") and post-column filters (e.g. the "GIF") represent different approaches which have their own advantages as well as experimental difficulties. This one day meeting is designed to explore the current state of the art, both with regard to the instrumentation and also applications of EFTEM in life and physical sciences.
Invited speakers: Dr Bernd Feja (Tietz Video & Image Processing Systems GbmH) Energy-filtered electron tomography Prof Joachim Mayer (Aachen University of Technology) EFTEM - the state of the art and future trends Dr Paul Midgley (University of Cambridge) EFTEM image series - taking elemental mapping into a new dimension Prof Michael Trendelenburg (German Cancer Research Center, DKFZ) EFTEM in biomedicine & biotechnology: Recent advances in specific element mapping
* CONTRIBUTIONS Contributed presentations are now being sought. Abstracts of no more than 200 words should be sent by email to: crispin.hetherington-at-materials.ox.ac.uk and should arrive by 31 May, 2001.
* REGISTRATION Registration will be Ł30 RMS/EMAG members, Ł15 students and Ł40 others and a printable form will be avialable from the new RMS website, from May on this page :
http://www.rms.org.uk/current%20events.html#eftem
Further details are available from the organisers: Dr Crispin Hetherington, tel. 01865 273799, crispin.hetherington-at-materials.ox.ac.uk Dr Laurence Tetley, tel. 0141 330 4431, l.tetley-at-bio.gla.ac.uk and from the meeting website : http://www-em.materials.ox.ac.uk/events/eftem.html
Note this meeting is timed to follow the 1-day FEGTEM III meeting to be held on 3 July 2001, also in Oxford (http://www-em.materials.ox.ac.uk/events/fegtem.html) **************************************************************************** ****************
Dr Laurence Tetley Division of Infection & Immunity, IBLS, Integrated Microscopy Facility Joseph Black Building University of Glasgow Glasgow G12 8QQ
********************* 1st Announcement and call for contributions *****************************
A meeting organised by the Royal Microscopical Society and supported by EMAG, FEI UK Ltd, JEOL (UK) Ltd.,LEO EM Inc., Gatan UK, TVIPS GmbH
DEVELOPMENTS IN ENERGY-FILTERED ELECTRON MICROSCOPY
Wednesday 4 July 2001
Department of Materials, Oxford University
The use of electron energy filters in analytical electron microscopy (EFTEM) is a relatively recent development that is proving to be an extremely powerful tool. In-column filters (e.g. so-called "omega-filters") and post-column filters (e.g. the "GIF") represent different approaches which have their own advantages as well as experimental difficulties. This one day meeting is designed to explore the current state of the art, both with regard to the instrumentation and also applications of EFTEM in life and physical sciences.
Invited speakers: Dr Bernd Feja (Tietz Video & Image Processing Systems GbmH) Energy-filtered electron tomography Prof Joachim Mayer (Aachen University of Technology) EFTEM - the state of the art and future trends Dr Paul Midgley (University of Cambridge) EFTEM image series - taking elemental mapping into a new dimension Prof Michael Trendelenburg (German Cancer Research Center, DKFZ) EFTEM in biomedicine & biotechnology: Recent advances in specific element mapping
* CONTRIBUTIONS Contributed presentations are now being sought. Abstracts of no more than 200 words should be sent by email to: crispin.hetherington-at-materials.ox.ac.uk and should arrive by 31 May, 2001.
* REGISTRATION Registration will be Ł30 RMS/EMAG members, Ł15 students and Ł40 others and a printable form will be avialable from the new RMS website, from May on this page :
http://www.rms.org.uk/current%20events.html#eftem
Further details are available from the organisers: Dr Crispin Hetherington, tel. 01865 273799, crispin.hetherington-at-materials.ox.ac.uk Dr Laurence Tetley, tel. 0141 330 4431, l.tetley-at-bio.gla.ac.uk and from the meeting website : http://www-em.materials.ox.ac.uk/events/eftem.html
Note this meeting is timed to follow the 1-day FEGTEM III meeting to be held on 3 July 2001, also in Oxford (http://www-em.materials.ox.ac.uk/events/fegtem.html) **************************************************************************** ****************
Dr Laurence Tetley Division of Infection & Immunity, IBLS, Integrated Microscopy Facility Joseph Black Building University of Glasgow Glasgow G12 8QQ
One thing that was not mentioned was the use of resin conditioners that help to preserve the knife edge of a glass knife. We use this with beginners. We have used lecithin and "PolyCut Ease" from Polysciences Information on this from a previous discussion can be found at http://www.biotech.ufl.edu/icbr/emcl/db/slippery.html I am not sure if a conditioner for LR White is mentioned there or in the Mollenhauer pub., but in my personal conversations with Hilton, he said that camphor could be used for acrylic resins. If this is of any interest I can dig deeper into this or perhaps we can contact Hilton Mollenhauer directly.
At 07:25 AM 4/26/2001 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Greg Erdos Assistant Director Biotechnology Program Ph. 352-392-1295 University of Florida Fax 352-846-0251 PO Box 118525 Gainesville, FL 32611 http://www.biotech.ufl.edu/~emcl
} If I'm not swamped, which is never, I do a two day processing. Using } a Lynx overnight to get into resin. I then put the specimens into molds and } let them cure for 24 hours. When I come in on day three I can cut thick } sections, have a conference with the Doc who submitted the case then cut } and stain the thins and then I'm ready to sit down at the scope.
} In reality, A specimen coming into our cutting room is submitted for } routine light microscopy and possibility special stains. Those sections are } then reviewed and the decision is made to submit the case for EM } examination. That can take one or two days. So there are four, possibility } five days before I can photograph the case. Another issue is } prioritization. I do live patients first and I do the tumors before } genetics cases. It makes no sense to work on a possible mitochondria } disorder when there is a kid with a chest wall tumor or a brain tumor. The } same goes for a kidney biopsy over a ciliary disorder. I have to prioritize } and frequently cases get put off a few days until the heats down. With a } current volume of about four hundred cases a year and an additional two } hundred cases for research I'm often asked "How long will this take?" My } usual answer which includes the scoping and printing or making a CD is five } working days. Autopsy's are not high on my list and periodically I'll hear } that the resident who submitted the case has rotated into another } department before the EM is done. To answer your question I'd have to know } what your case load is and how many skilled hands are available.
} Howard Mulhern } Supv. Path. EM Facility } Children's Hospital } Dept. of Pathology } 300 Longwood Ave. } Boston, 02115 Ma. } } } }
I have been using a Chien grid to pick up sections, like a loop, and place it on top of a coated grid. Let water evaporate and sections will drop on to the grid below. I dip Chien grids in 1% formvar, shake off excess and place then on a filter paper to dry. This treatment ensures water film is maintained during the transfer. Cleaning the Chien grids in acid has the same effect. The handle of Chien grid is bended for better handling. I have tried a "perfect loop" and I prefer Chien grids.
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
} } } Claudia Hayward-Costa {LS_S562-at-crystal.kingston.ac.uk} 04/25 5:57 AM } } } ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear Microscopists,
I wondered how commonly people use a tool (like a loop) to pick up ultrathin sections.
Does it make life easier or does it only introduce different problems?
I was told that I can buy some "luxury" - but am still undecided.
Your views would be very much appreciated.
Regards
Claudia
Dr. C. Hayward-Costa School of Life Sciences Kingston University Penrhyn Road, Kingston upon Thames Surrey KT1 2EE, UK 44(0)208 547 2000 x 2240 Email: c.hayward-at-kingston.ac.uk Fax: 44(0)208 547 7562
} -----Original Message----- } From: Earl Weltmer [SMTP:eweltmer-at-home.com] } Sent: Wednesday, April 25, 2001 8:19 PM } To: Chuck Butterick; Microscopy-at-sparc5.microscopy.com } Subject: Re: Apology } } ------------------------------------------------------------------------ } 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 rather enjoyed it. } } Earl } } ----- Original Message ----- } } From: "Chuck Butterick" {cbutte-at-ameripol.com} } To: {Microscopy-at-sparc5.microscopy.com} } Sent: Wednesday, April 25, 2001 5:36 AM } Subject: Apology } } } } ------------------------------------------------------------------------ } } 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 } } -----------------------------------------------------------------------. } } } } } } Listers, } } } } A slip of the finger caused me to send an un-topical joke to all. } } While no has yet criticized (though some have expressed } appreciation } } and/or agreement), I feel that the joke was inappropriate to the } } venue. My apologies. } } } } Chuck Butterick } } } } }
OK this is going to seem like a stupid question. I am going to try araldite 502 embedding media. the instruction sheet calls for a volumetric measurement. I would prefer to measure the each chemical in a balance. OK here is the question: do i multiply the specific gravity by the volume or divide it. it has been a long time for me since chemistry 101. thanks
Dear J, At the beginning of every semester I give three general hints on calculations to my students. The relevant one for you is always write down units explicitly. Since the density (numerically equal to the specific gravity in cgs units) is in g/cm^3, if you multiply the volume (in cm^3) by the density, you get the mass. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
Following the recent discussion thread of scanners, I wanted to say that we are the proud owners of a Polaroid SprintScan 45 Ultra, and are very happy with the results we have gotten from it. It came with a number of negative holders covering a range of sizes from 35mm to 4x5, except one for 3.25x4.25 TEM sheet film. Right now we're trying to think of ways around this problem and would be happy to hear offline from anyone who has some ideas and/or solutions. I would even entertain the idea of having a holder made (this is not an option locally for me).
Thanks in advance for any help (and there is usually plenty, thanks to this list!).
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
Paula Allan-Wojtas Research Scientist - Food Microstructure Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
Here in the lab we still do the hand processing.. not automated processor yet.
There are two techs, but turnaround time is different between both of us.. i.e. one of us works faster than the other...
Normally when a biopsy comes in on say Monday.. I can have images by Wednesday.. Out here we do not use a darkroom anymore.. Our lab has gone digital with a AMT system.
The Pathologists are thrilled with it since it cuts down our turnaround time from 5 days to 3 days...
Does anyone knows where I can find a five-axes (four-axes U-Stage is fine too) universal stage for sale or donation?
Thank all of you in advance.
Best wishes,
Leonardo -- --- Leonardo Lagoeiro Departamento de Geologia Universidade Federal de Ouro Preto Ouro Preto, MG, 35400-000 Brazil E-mai: lagoeiro-at-degeo.ufop.br
I would like to start up a thread about the level of instruction and the amount of time that undergraduates spend in an introductory EM course that covers TEM & SEM. The need comes from low numbers of student credit hours generated verses the amount of time spent by the students and instructor in the course. For example, I usually have between 2 to 4 students in my four credit hour course, taught every fall sememster. The students receive two one hour lectures per week, two supervised two hour labs and a third unsupervised two hour lab. I teach the students in pairs during the intensive hands-on portions like ultramicrotomy and scope operations, so I usually have two lab sections to prep for, or about eight contact hours for labs per week. With the lectures, I then have ten hours of contact per week for the course.
What are the teaching loads and class sizes of other instructors who teach undergrad EM?
Thanks
Robert
Robert Fitton Teaching Associate/Director of Laboratories Luther College Department of Biology 700 College Drive Decorah, IA 52101
Voice 563-387-1559 FAX 563-387-1080 (If the 563 area code does not work, try our old area code of 319)
Enjoy a visit to our website: http://www.luther.edu/~biodept/
Our TAT differs depending on the service rendered.
Diagnostic virology: 15 min to ~2 hours (depending on whether we ultracentrifuge) for negative staining of bodily fluids. 2-3 days for thin sections of tissue (depending on how many blocks we have to cut and how long we have to look if the sample shows no viruses before we call it negative). Three super techs do ~700 negative stain specimens and 100 thin section samples of clinical material per year, plus a number of research samples (20-40, some of which are cryosections for IEM).
Surgical pathology: 2 days (about 32 hours). In on a morning, into the oven that night, cut, photographed, and dark room micrographs printed and delivered to the pathologist by 5 PM the next day. (We're working on going digital.) Two super techs do ~500 clinical samples plus ~100 research samples a year.
Five techs total. They can cross cover, but each has a specialty, and generally, the same 3 do virology, and the other 2 do surgical pathology.
Sara E. Miller, Ph. D. P. O. Box 3712 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-3265
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} Dear Microscopist,
} Does anyone knows where I can find a five-axes (four-axes U-Stage is } fine too) universal stage for sale or donation?
} Thank all of you in advance.
} Best wishes,
} Leonardo } -- } --- } Leonardo Lagoeiro } Departamento de Geologia } Universidade Federal de Ouro Preto } Ouro Preto, MG, 35400-000 } Brazil } E-mai: lagoeiro-at-degeo.ufop.br
The message you sent to jeolbxl-at-pophost.eunet.be could not be delivered. Either you misspelled your correspondent's name, or jeolbxl no longer exists in the pophost.eunet.be domain.
If you want more information, you can contact the postmaster at postmaster-at-pophost.eunet.be. Please understand that we can not give emailaddresses from our customers.
Kind Regards,
The mail delivery system. -- --- Leonardo Lagoeiro Departamento de Geologia Universidade Federal de Ouro Preto Ouro Preto, MG, 35400-000 Brazil E-mai: lagoeiro-at-degeo.ufop.br
Polaroid has a small sheet insert that fits into the 4 X 5 and holds the negative down with small magnets. You need to call someone at Polaroid and request this. They gave them to me and a few others on the List for free. I originally talked to John Warren WARRENJ1-at-POLAROID.COM There is also a Dennis Lizier, but I do not have his contact information. -Scott
-----Original Message----- } From: Paula Allan-Wojtas [mailto:AllanWojtasP-at-em.agr.ca] Sent: Thursday, April 26, 2001 12:17 PM To: microscopy-at-sparc5.microscopy.com
Hi, all,
Following the recent discussion thread of scanners, I wanted to say that we are the proud owners of a Polaroid SprintScan 45 Ultra, and are very happy with the results we have gotten from it. It came with a number of negative holders covering a range of sizes from 35mm to 4x5, except one for 3.25x4.25 TEM sheet film. Right now we're trying to think of ways around this problem and would be happy to hear offline from anyone who has some ideas and/or solutions. I would even entertain the idea of having a holder made (this is not an option locally for me).
Thanks in advance for any help (and there is usually plenty, thanks to this list!).
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
Paula Allan-Wojtas Research Scientist - Food Microstructure Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
Robert Any undergraduate teaching of EM techniques we do strictly within the context of student projects, chiefly in the final honours year (4th year of the BSc course at Edinburgh). Since I joined the Botany department staff in 1976 the policy has been not to teach methods. I don't especially agree with that policy, but it is the way things have always been done here. I get the luxury of a 1-hour presentation on EM to first year biology students at the end of their first year. We also demonstrate the EMs to class groups from various departments, and we train a few students per year as they require access to EM for project work, but there have been no formal taught undergraduate courses with hands-on practical tuition in EM in the Science Faculty at Edinburgh for many years now. Chris
----- Original Message ----- } From: "Robert Fitton" {fittonro-at-luther.edu} To: {Microscopy-at-sparc5.microscopy.com} Sent: Thursday, April 26, 2001 9:40 PM
FW: Re: Film Processing and dynamic range and Exposure of film to electrons
} A couple of thoughts on film: } Kodak special publication P-116 describes the interaction of electrons with film (from the point of view of people that produce the film-I assume they study it fairly closely): } "Each electron in an incident beam may pass through perhaps 30 or more silver halide grains before it is stopped completely...........The trajectory is a changing one as collisions occur, and energy is lost to the silver halide grains, and to a lesser extent, to the gelatin in which the silver halide grains are dispersed. This transfer of energy, which is low at the beginning of an electron path and increases as the electron is slowed by collisions, is responsible for the formation of specs of silver atoms in the affected grain. If the aggregate of silver atoms formed is sufficiently large-believed to be between 3 and 6 atoms, the entire grain will be capable of being converted to metallic silver during photographic development. While the passage of a single electron may not render each grain developable, to which it gives up energy, overall sensitivity is such that normally at least one grain, and very likely, a number of grains, will be converted to metallic silver. If this is the case, the photographic material records all of the information in the electron beam;" } This document further discusses contrast, illustrating the sensitivity of the film with log exposure/density curves, but only up to a density of 2. It may be worth noting that John Spence states in the appendix of his text that for photographic emulsions, the exposure/density curve is linear up to a density of between one and 2. The departure from linearity would seem to be related to some sort of "pile-up" effect. While the exposure/density curves may be linear over this range, the read-out of the film is logarithmic, as the density is defined as the log base 10 of the transmitted light intensity divided by the incident light intensity. That is why, for example, one commonly used, and successful method of increasing contrast is to increase the exposure level of the film. For example, a 10% change in electron dose near a density of 1 will produce a 25% change in transmitted light intensity, where a 10 % change in dose near a density of 2 will produce a 58% change in transmitted light intensity. This logarithmic read-out characteristic is perhaps why all of the Kodak sensitivity plots are in log dose/density format, and they are curves. Kodak also illustrates how the contrast of the print can be estimated from the slope of these log dose/density curves, since the slope increases with average density. In summary, even though the density of the film may be linear with electron dose, what you see in a print, what you get from a scanner, even what you see from observing the film on a light box, is logarithmic with respect to electron dose. This is also why images obtained with a CCD camera or an image plate, which are linear with electron dose, have a distinctly different appearance than images obtained with film unless digital gamma is used to adjust the display. So, everyone was essentially correct in what they said, it's just that they were not describing the same thing. } One may want to look at the claims made for Fuji image plates on their web site, which can be accessed at "fujimed.com/sub/FDL5000.pdf". They show plots of log readout intensity/log dose for the Fuji image plate superimposed on density/log dose for film. The Fuji image plate is more linear than film, but for the range 0-2, film is relatively parallel to the image plate. Beyond a density of 2, the nonlinearity becomes gradually more pronounced, but there is never any saturation encountered. } Finally, I would like to reiterate that this has been a fascinating string, and it seems that everyone has contributed some useful and thought provoking information.} } } John Mardinly } Intel Materials Technology
I have the pleasure of teaching full semester (13 week) EM course to undergraduates at Queensland University of Technology in Brisbane, Australia. We've offered this course for quite a number of years - in fact, even had it extended from part of a broader topic to a full semester just for EM! The course is offered as part of the School of Life Sciences undergraduate program and, hence, primarily addresses biological EM. However, it is open to students from other disciplines - presently we do not have a full EM course in the physical sciences, although my colleague Dr Thor Bostrom teaches several weeks of EM in various courses for physics, chemistry and engineering undergraduate students.
Apart from this, I will usually run single 2 hour practical sessions for a couple of other undergraduate courses (mainly microbiology, where students look at negatively stained microbes they have isolated from various sources). I'll often be asked to give a half hour (sometimes an hour) introductory EM lecture to these students, prior to the practicals.
I also teach a brief introduction (2-3 weeks) to EM as part of a post-graduate course on cell structure and function in pathology. Other post-graduates needing EM as part of their research are taught individually by staff in our EM Facility, to the level that their projects require.
My full semester EM course is offered in the third year of the B. Applied Science course. This year I have 30 students - the numbers have been increasing over the past few years from a previous average of 20. I usually also have a couple of post-graduate (research) students attending the lectures - these students will be using our EM facilities as part of their research, and I find this is an effective means of giving them some useful background and theory to support their laboratory work.
For this course, I am allocated two by one hour lectures each week (total of 26 hours lectures for the semester). I cover basic instrumentation (TEM, SEM, preparation equipment) and biological sample preparation methods, including cryotechniques. I also include immunolabelling methods, some cytochemistry, microanalysis (Thor Bostrom gives these lectures), a few interesting "non-biological" applications, and a brief introduction to some other imaging methods (confocal, AFM etc).
Practical sessions have been a bit of a "challenge" as student numbers have increased and EM staff numbers have decreased! However, I've now got a system that works, and doesn't cause me (or the equipment) too much stress. I have two by 3 hours practical sessions running each week - ie. 6 hours of my teaching time per week (total of 72 hours of my teaching time for the semester).
Each student attends a total of 24 hours practical sessions for the semester (this fits with our "official" allocation for total practical time for a course of this credit point value). I divide the class into small groups - 3 to 4 students per group, with current student numbers. For preparation lab based pracs (where we have enough space and the students don't need such close supervision), I'll have a number of these small groups attending the same practical session. For EM operation and viewing pracs, I'll only have one small group attending at a time (usually 1 or 1.5 hours of the session for each small group). This has enabled me to run practical sessions almost single-handed - money to employ tutors/demonstrators or other teaching assistants has been a bit difficult to obtain! If anyone would like further information on the practicals I run or on the logistics of this, please contact me by direct email - does anyone else out there run EM practicals for undergraduates??? I haven't seen a rush of emails in response to Robert's questions.
Fortunately, my university is still keen to have students doing "hands-on" practicals in their courses. However, I've certainly had to decrease practical work substantially since I first started teaching in this EM course. A lot of this is due to decreased staff numbers in our EM Facility - a problem that I guess most of us face. I teach other courses (microbiology, parasitology) and encounter the same problems there - it is now very difficult to get competent, experienced people to assist in practical classes, particularly for more advanced or specialised courses.
On the bright side, I've found the students enjoy EM a great deal - and it is one of the few opportunities our undergraduates get to use expensive and (relatively!) modern instrumentation, even if it is only on a very limited scale. I've not had many students who don't show a great deal of enthusiasm when told it is now their turn to operate the TEM or SEM and take some micrographs. This semester, more so than previously, at the end of each practical session quite a number of my students have taken the time to thank me for running the practicals (even after the night classes which finish at 8pm). I figure that has got to be a good sign!!!
Regards Deb ***************************************************** Dr Deborah Stenzel Lecturer (Microbiology) School of Life Sciences and Applications Specialist (Biological) Analytical Electron Microscopy Facility Queensland University of Technology GPO Box 2434 Brisbane 4001 Australia
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Hi, everyone, Now, I need to simulate HRTEM image of a phase that I can not find its space group in X-ray powder diffraction files. Do you know whether it is possible for me to find its atomic positions. Could you please give me suggestions? Thank you very much in advance. WY
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Dear Listers, We are in need to prepare a short course on EM basics. I would be thankfull for any information about free internet resources or commercially availiable CDs containing illustative materials, animations, programms illustrating image formation (not simulating software) that we can use in the lectures and during the seminars.
EuroFE 2001, the European Meeting on Applications of Field Emission Technologies, EuroFE 2001 will tale palace this year in University of Alicante, Spain from November 12-16, 2001. Details are available at www.cmp-cientifica.com/eurofe2001
The fundamental purpose of workshops such as EUROFE2001 consists in bringing together a hundred scientific, leaders in this area, in order to: Link all active research groups in Field Emission.
Link all industries interested in Field Emission. At EUROFE2001, major companies such as Motorola (USA), Candescent (USA), Samsung (Korea), PixTech (France), Saint-Gobain (France), Thomson (France), LG (Korea) and PFE (UK) will be present.
Foster co-operation and the interchange of ideas between research groups and European Industry.
Provide an overview of the research and commercialisation of Field Emission technologies within Europe. During EUROFE2001, a session will be dedicated to Space Applications (ESA/ESTEC speakers).
Allow rapid and flexible response to new technological challenges
Based on the format established at EUROFE2000, ample time for discussion will be available for meaningful interaction between senior researchers/graduate students and industrial partners.
Funds to offset travel expenses for graduate students to present their work in a poster session at EUROFE2001 will be available.
For further details please contact:
Antonio Correia CMP Cientifica Apdo. Correos 20 28230 Las Rozas (Madrid), Spain Tel: +34 91 6407187 Fax: +34 91 6407186
Regards,
Tim
***************************************************************** Tim E. Harper CEO CMP Cientifica s.l. Space & NanoTechnology Division Phone +34 91 640 71 85 Fax +34 91 640 71 86 http://www.cmp-cientifica.com/
Hi, everyone, Now, I need to simulate HRTEM image of a phase that I can not find its space group in X-ray powder diffraction files. Do you know whether it is possible for me to find its atomic positions. Could you please give me suggestions? Thank you very much in advance. WY
Carbon black in polymer or rubber will cause no more problems than most any other hydrocarbon. The electron beam does sublimate the plastic. Especially when working with pure carbon black, one can essentially watch a contaminant layer grow on the constituent particles of a given aggregate. A cold finger is a very good idea but not absolutely required if you are just scanning the sample.
Our Philips 300 is still in excellent operating condition after 28 years of carbon black work.
Chuck Butterick Engineered Carbons, Inc. Borger, TX
by ultra5.microscopy.com (8.9.3+Sun/8.9.1) id HAA26160 for dist-Microscopy; Fri, 27 Apr 2001 07:37:15 -0500 (CDT) Received: from no_more_spam.com (sparc5 [206.69.208.10]) by ultra5.microscopy.com (8.9.3+Sun/8.9.1) with SMTP id HAA26157 for "MicroscopyFilteredEmail1-at-msa.microscopy.com"; Fri, 27 Apr 2001 07:36:45 -0500 (CDT) Received: from mercury.uwe.ac.uk (mercury.uwe.ac.uk [164.11.132.23]) by ultra5.microscopy.com (8.9.3+Sun/8.9.1) with ESMTP id HAA26150 for {Microscopy-at-sparc5.microscopy.com} ; Fri, 27 Apr 2001 07:36:33 -0500 (CDT) Received: from fas655.uwe.ac.uk ([164.11.205.145]) by mercury.uwe.ac.uk (2.0.4/SMS 2.0.4-devel) with SMTP id NAA04426; Fri, 27 Apr 2001 13:33:48 +0100 (BST)
Back in 1989 all Biology degree and sub-degree students attended 3 sessions in the EM Unit. Now the expansion in numbers precludes such demonstrations. I regret this change.
In the first year they get a few lectures and a "dry" class identifying cell organelles. In the final year about 6 students do major projects in the lab.
We still do undergraduate classes in dust ID and in EDX for modules taken by Chemistry, Environmental Science and Environmental Health students as the numbers are smaller.
Dave
On Fri, 27 Apr 2001 16:08:08 +1000 Dr Deborah Stenzel {d.stenzel-at-qut.edu.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 Robert and all } } I have the pleasure of teaching full semester (13 week) EM course to } undergraduates at Queensland University of Technology in Brisbane, } Australia. We've offered this course for quite a number of years - in } fact, even had it extended from part of a broader topic to a full semester } just for EM! The course is offered as part of the School of Life Sciences } undergraduate program and, hence, primarily addresses biological } EM. However, it is open to students from other disciplines - presently we } do not have a full EM course in the physical sciences, although my } colleague Dr Thor Bostrom teaches several weeks of EM in various courses } for physics, chemistry and engineering undergraduate students. } } Apart from this, I will usually run single 2 hour practical sessions for a } couple of other undergraduate courses (mainly microbiology, where students } look at negatively stained microbes they have isolated from various } sources). I'll often be asked to give a half hour (sometimes an hour) } introductory EM lecture to these students, prior to the practicals. } } I also teach a brief introduction (2-3 weeks) to EM as part of a } post-graduate course on cell structure and function in pathology. Other } post-graduates needing EM as part of their research are taught individually } by staff in our EM Facility, to the level that their projects require. } } } My full semester EM course is offered in the third year of the B. Applied } Science course. This year I have 30 students - the numbers have been } increasing over the past few years from a previous average of 20. I } usually also have a couple of post-graduate (research) students attending } the lectures - these students will be using our EM facilities as part of } their research, and I find this is an effective means of giving them some } useful background and theory to support their laboratory work. } } For this course, I am allocated two by one hour lectures each week (total } of 26 hours lectures for the semester). I cover basic instrumentation } (TEM, SEM, preparation equipment) and biological sample preparation } methods, including cryotechniques. I also include immunolabelling methods, } some cytochemistry, microanalysis (Thor Bostrom gives these lectures), a } few interesting "non-biological" applications, and a brief introduction to } some other imaging methods (confocal, AFM etc). } } Practical sessions have been a bit of a "challenge" as student numbers have } increased and EM staff numbers have decreased! However, I've now got a } system that works, and doesn't cause me (or the equipment) too much } stress. I have two by 3 hours practical sessions running each week - ie. 6 } hours of my teaching time per week (total of 72 hours of my teaching time } for the semester). } } Each student attends a total of 24 hours practical sessions for the } semester (this fits with our "official" allocation for total practical time } for a course of this credit point value). I divide the class into small } groups - 3 to 4 students per group, with current student numbers. For } preparation lab based pracs (where we have enough space and the students } don't need such close supervision), I'll have a number of these small } groups attending the same practical session. For EM operation and viewing } pracs, I'll only have one small group attending at a time (usually 1 or 1.5 } hours of the session for each small group). This has enabled me to run } practical sessions almost single-handed - money to employ } tutors/demonstrators or other teaching assistants has been a bit difficult } to obtain! If anyone would like further information on the practicals I } run or on the logistics of this, please contact me by direct email - does } anyone else out there run EM practicals for undergraduates??? I haven't } seen a rush of emails in response to Robert's questions. } } Fortunately, my university is still keen to have students doing "hands-on" } practicals in their courses. However, I've certainly had to decrease } practical work substantially since I first started teaching in this EM } course. A lot of this is due to decreased staff numbers in our EM Facility } - a problem that I guess most of us face. I teach other courses } (microbiology, parasitology) and encounter the same problems there - it is } now very difficult to get competent, experienced people to assist in } practical classes, particularly for more advanced or specialised courses. } } On the bright side, I've found the students enjoy EM a great deal - and it } is one of the few opportunities our undergraduates get to use expensive and } (relatively!) modern instrumentation, even if it is only on a very limited } scale. I've not had many students who don't show a great deal of } enthusiasm when told it is now their turn to operate the TEM or SEM and } take some micrographs. This semester, more so than previously, at the end } of each practical session quite a number of my students have taken the time } to thank me for running the practicals (even after the night classes which } finish at 8pm). I figure that has got to be a good sign!!! } } } Regards } Deb } ***************************************************** } Dr Deborah Stenzel } Lecturer (Microbiology) } School of Life Sciences } and } Applications Specialist (Biological) } Analytical Electron Microscopy Facility } Queensland University of Technology } GPO Box 2434 } Brisbane 4001 } Australia } } Phone + 61 7 3864 5036 } Fax + 61 7 3864 5100 } email d.stenzel-at-qut.edu.au } } http://www.sci.qut.edu.au/aemf } }
---------------------------------------- Patton, David Email: David.Patton-at-uwe.ac.uk "University of the West of England"
You make mention of "the EM staff" - I'm wondering how many people are actually involved in teaching the course components, and how much time do they collectively spend, for example per student per week??
I'm curious to know because we have two people involved in teaching a full semester Bio EM course for up to 8 students. The students in the end turn in a fairly polished portfolio (with about a dozen 8x10 micrographs plus figure legends, from three different tissues that they have processed, cut, and scoped).
Questions have recently arisen as to how many man-hours we 'should' allot to such a course, which is far outside the norm for our other lab courses.
I'm interested in off-line dialogue, if others are.
Ann Hein Lehman EM Facility Manager Trinity College Hartford CT 06106 v. 860-297-4289 e. ann.lehman-at-trincoll.edu
-----Original Message----- } From: Dr Deborah Stenzel [mailto:d.stenzel-at-qut.edu.au] Sent: Friday, April 27, 2001 2:08 AM To: Microscopy-at-sparc5.microscopy.com
Dear Robert and all
I have the pleasure of teaching full semester (13 week) EM course to undergraduates at Queensland University of Technology in Brisbane, Australia. We've offered this course for quite a number of years - in fact, even had it extended from part of a broader topic to a full semester just for EM! The course is offered as part of the School of Life Sciences undergraduate program and, hence, primarily addresses biological EM. However, it is open to students from other disciplines - presently we do not have a full EM course in the physical sciences, although my colleague Dr Thor Bostrom teaches several weeks of EM in various courses for physics, chemistry and engineering undergraduate students.
Apart from this, I will usually run single 2 hour practical sessions for a couple of other undergraduate courses (mainly microbiology, where students look at negatively stained microbes they have isolated from various sources). I'll often be asked to give a half hour (sometimes an hour) introductory EM lecture to these students, prior to the practicals.
I also teach a brief introduction (2-3 weeks) to EM as part of a post-graduate course on cell structure and function in pathology. Other post-graduates needing EM as part of their research are taught individually by staff in our EM Facility, to the level that their projects require.
My full semester EM course is offered in the third year of the B. Applied Science course. This year I have 30 students - the numbers have been increasing over the past few years from a previous average of 20. I usually also have a couple of post-graduate (research) students attending the lectures - these students will be using our EM facilities as part of their research, and I find this is an effective means of giving them some useful background and theory to support their laboratory work.
For this course, I am allocated two by one hour lectures each week (total of 26 hours lectures for the semester). I cover basic instrumentation (TEM, SEM, preparation equipment) and biological sample preparation methods, including cryotechniques. I also include immunolabelling methods, some cytochemistry, microanalysis (Thor Bostrom gives these lectures), a few interesting "non-biological" applications, and a brief introduction to some other imaging methods (confocal, AFM etc).
Practical sessions have been a bit of a "challenge" as student numbers have increased and EM staff numbers have decreased! However, I've now got a system that works, and doesn't cause me (or the equipment) too much stress. I have two by 3 hours practical sessions running each week - ie. 6 hours of my teaching time per week (total of 72 hours of my teaching time for the semester).
Each student attends a total of 24 hours practical sessions for the semester (this fits with our "official" allocation for total practical time for a course of this credit point value). I divide the class into small groups - 3 to 4 students per group, with current student numbers. For preparation lab based pracs (where we have enough space and the students don't need such close supervision), I'll have a number of these small groups attending the same practical session. For EM operation and viewing pracs, I'll only have one small group attending at a time (usually 1 or 1.5 hours of the session for each small group). This has enabled me to run practical sessions almost single-handed - money to employ tutors/demonstrators or other teaching assistants has been a bit difficult to obtain! If anyone would like further information on the practicals I run or on the logistics of this, please contact me by direct email - does anyone else out there run EM practicals for undergraduates??? I haven't seen a rush of emails in response to Robert's questions.
Fortunately, my university is still keen to have students doing "hands-on" practicals in their courses. However, I've certainly had to decrease practical work substantially since I first started teaching in this EM course. A lot of this is due to decreased staff numbers in our EM Facility - a problem that I guess most of us face. I teach other courses (microbiology, parasitology) and encounter the same problems there - it is now very difficult to get competent, experienced people to assist in practical classes, particularly for more advanced or specialised courses.
On the bright side, I've found the students enjoy EM a great deal - and it is one of the few opportunities our undergraduates get to use expensive and (relatively!) modern instrumentation, even if it is only on a very limited scale. I've not had many students who don't show a great deal of enthusiasm when told it is now their turn to operate the TEM or SEM and take some micrographs. This semester, more so than previously, at the end of each practical session quite a number of my students have taken the time to thank me for running the practicals (even after the night classes which finish at 8pm). I figure that has got to be a good sign!!!
Regards Deb ***************************************************** Dr Deborah Stenzel Lecturer (Microbiology) School of Life Sciences and Applications Specialist (Biological) Analytical Electron Microscopy Facility Queensland University of Technology GPO Box 2434 Brisbane 4001 Australia
Like Chris, our department does not teach techniques courses. However, I include SEM and TEM in a course called Microanatomy (about 16 students every fall semester). What I have done is combine more-or-less traditional histology class lectures with a lab component in which students compare the histology of the same organ from a mouse and frog. They make their own paraffin, methacrylate, and epoxy plastic sections and critical point dry their other samples. I spend a great deal of time on light microscopy theory, how to clean and align a light microscope, capture digital images, and prepare micrographs for illustration (scale bars and labels). I do not spend a lot of time on the use of the electron microscopes apart from showing students enough for them to change mag, focus, and take a picture. Since I can't do everything, I've decided that more students will find basic light microscopy more generally useful in the future than EM techniques. However, the students really love using the "big iron" and are especially enthusiastic about SEM.
Cutting thin sections with glass knives has also been a bottleneck for my students so I was glad to see the discussion on this recently. Thanks, Dick, for summarizing the responses. -- Gary P. Radice gradice-at-richmond.edu Associate Professor of Biology 804 289 8107 (voice) University of Richmond 804 289 8233 (FAX) Richmond VA 23173 http://www.science.richmond.edu/~radice
We took one of the smaller negative holders for which we didn't foresee a need and submitted it along with a TEM negative to a machinist in our shop. He was able to mill the opening such that it was the ideal size for the 3.25 x 4.25 negative. To give it that professional look he completed the job by touching up the milled edges with black paint.
Paul
Paul J. Gerroir Microscopy Materials Characterization Xerox Research Centre of Canada 2660 Speakman Drive Mississauga, Ontario L5K 2L1
-----Original Message----- } From: Paula Allan-Wojtas [mailto:AllanWojtasP-at-em.agr.ca] Sent: Thursday, April 26, 2001 12:17 PM To: microscopy-at-sparc5.microscopy.com
Hi, all,
Following the recent discussion thread of scanners, I wanted to say that we are the proud owners of a Polaroid SprintScan 45 Ultra, and are very happy with the results we have gotten from it. It came with a number of negative holders covering a range of sizes from 35mm to 4x5, except one for 3.25x4.25 TEM sheet film. Right now we're trying to think of ways around this problem and would be happy to hear offline from anyone who has some ideas and/or solutions. I would even entertain the idea of having a holder made (this is not an option locally for me).
Thanks in advance for any help (and there is usually plenty, thanks to this list!).
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
Paula Allan-Wojtas Research Scientist - Food Microstructure Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
Hello, I feel that I must add to this because as a general statement I believe it is mis-leading ... with respect to carbon black & many other carbon species... some 7000 images of carbon later I suggest that watching carbon morph in the beam is an exception not the rule. If you sit on anything long enough it will collect carbon but let's remain objective. Among other things, I have looked at CBs right out of reactors & extracted from polymers. How clean your instrument is can be a significant issue. My tool is a JEOL 2010, LaB6 operating at ~2x10-5Pa, & I use an LN2 cooled anti contamination device, ACD in Jeol lingo. The stage is room temperature and I usually work at 100 kV for the contrast enhancement. I have witnessed such things in experimental materials that I will term "highly reactive carbons". This lore of morphing carbon may be the most commonly used challenge of conclusions drawn from TEM data. Stories of exceptions tend to propagate over space & time. Don't get me wrong, it does happen.
Diverging a bit...
Of much more concern to anyone working in carbon is the carbon contamination supplied on new, right out of the box carbon coated grid formvar &/or Lacey carbon grid. The is a real problem for the manufactures & to my knowledge, despite any claims you may hear, no one has solved this problem. Once you understand & know the many habits carbon can have, that is the stuff supplied on new grids, you will realize that many images you see in the literature could easily be store bought carbon grid contamination. I have personally recorded dozens of such images... onions, fibers, rods, angular faceted bits, plates, turbostratic plates, polycrystalline graphite, CB looking stuff, graphitic ribbons & oh the nano structures, horns, fibers, .... Oh yea, watch out for very thin, low contrast crystals that throw a textbook perfect [001] hexagonal DP. it is not carbon. I don't know what it is. Weather on not you have realized it, you have seen them in imaging mode & probably passed them off as film thickness irregularities. I am looking at an image computer directory titled "grid stuff". it contains 12 images recorded on one new out of the box lacey carbon grid. Their titles are: hex plate, nest-2, nest -1, plates, rapping ribbon, plates plus, ribbon, fiber, rodlike, nanohorn, elongated hex. To all of this, let me refer you to an article recently or soon to be published in Carbon. I believe the author's name is Harris. I apologize for the poor quality reference, I've spaced out my copy of the preprint. It pretty much reads as something I have contemplated writing over the years but...
it's real folks.
Bruce Brinson Rice U.
Guess there is a reason ASTM standards require 1000 representative images.
Chuck Butterick 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 } -----------------------------------------------------------------------. } } Carbon black in polymer or rubber will cause no more problems than } most any other hydrocarbon. The electron beam does sublimate the } plastic. Especially when working with pure carbon black, one can } essentially watch a contaminant layer grow on the constituent } particles of a given aggregate. A cold finger is a very good idea but } not absolutely required if you are just scanning the sample. } } Our Philips 300 is still in excellent operating condition after 28 } years of carbon black work. } } Chuck Butterick } Engineered Carbons, Inc. } Borger, TX } } ______________________________ Reply Separator _________________________________ } Subject: carbon black in polymer } Author: Margaret Miller {MILLERMM-at-uthscsa.edu} at INTERNET-MAIL } Date: 4/26/01 2:27 PM } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Does carbon black cause a problem in the TEM? What percautions should I } take? Is the use of the cold finger recommended? } }
Just turn the So-163 film sideways on the 4x5 holder.
Paul PPG Industries
-----Original Message----- } From: Gerroir, Paul J [mailto:Paul.Gerroir-at-crt.xerox.com] Sent: Friday, April 27, 2001 6:28 AM To: Paula Allan-Wojtas; microscopy-at-sparc5.microscopy.com
Paula,
We took one of the smaller negative holders for which we didn't foresee a need and submitted it along with a TEM negative to a machinist in our shop. He was able to mill the opening such that it was the ideal size for the 3.25 x 4.25 negative. To give it that professional look he completed the job by touching up the milled edges with black paint.
Paul
Paul J. Gerroir Microscopy Materials Characterization Xerox Research Centre of Canada 2660 Speakman Drive Mississauga, Ontario L5K 2L1
-----Original Message----- } From: Paula Allan-Wojtas [mailto:AllanWojtasP-at-em.agr.ca] Sent: Thursday, April 26, 2001 12:17 PM To: microscopy-at-sparc5.microscopy.com
Hi, all,
Following the recent discussion thread of scanners, I wanted to say that we are the proud owners of a Polaroid SprintScan 45 Ultra, and are very happy with the results we have gotten from it. It came with a number of negative holders covering a range of sizes from 35mm to 4x5, except one for 3.25x4.25 TEM sheet film. Right now we're trying to think of ways around this problem and would be happy to hear offline from anyone who has some ideas and/or solutions. I would even entertain the idea of having a holder made (this is not an option locally for me).
Thanks in advance for any help (and there is usually plenty, thanks to this list!).
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
Paula Allan-Wojtas Research Scientist - Food Microstructure Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
I used to teach an EM course to the graduate students here. It would run for 10 weeks and would consist of a 2-2.5 hour lecture period followed by a lunch break and then a 4 hour lab session during which I would teach them the "technique du jour" (fixation, embedding, block trimming, thick sectioning, thin sectioning, SEM sample prep, darkroom techniques, use of the EM....and in my really crazy days, alignment of the column!). the class would then be expected to come in during the week to put into practice what we had done in class. by the end of the class I expected each to have produces block, slides with stained thicks, grids with thins and 8X10 printed micrographs that showed me more than highly enlarged organelles.
I was also supposed to be running the dept's EM facility through all this. I would tell all of my usual clients not to expect anything substantial from me while the course ran, because I was too busy holding the hands of my 5-10 students. It would amount to 20-30 contact hours/week for the 10 week run. Exhausting.
Now that my facility is a college core, I can't just "drop out" for 10 weeks, and so the course is no longer offered.
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 Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear Deb and others,
An interesting thread...
You make mention of "the EM staff" - I'm wondering how many people are actually involved in teaching the course components, and how much time do they collectively spend, for example per student per week??
I'm curious to know because we have two people involved in teaching a full semester Bio EM course for up to 8 students. The students in the end turn in a fairly polished portfolio (with about a dozen 8x10 micrographs plus figure legends, from three different tissues that they have processed, cut, and scoped).
Questions have recently arisen as to how many man-hours we 'should' allot to such a course, which is far outside the norm for our other lab courses.
I'm interested in off-line dialogue, if others are.
Ann Hein Lehman EM Facility Manager Trinity College Hartford CT 06106 v. 860-297-4289 e. ann.lehman-at-trincoll.edu
-----Original Message----- } From: Dr Deborah Stenzel [mailto:d.stenzel-at-qut.edu.au] Sent: Friday, April 27, 2001 2:08 AM To: Microscopy-at-sparc5.microscopy.com
Dear Robert and all
I have the pleasure of teaching full semester (13 week) EM course to undergraduates at Queensland University of Technology in Brisbane, Australia. We've offered this course for quite a number of years - in fact, even had it extended from part of a broader topic to a full semester just for EM! The course is offered as part of the School of Life Sciences undergraduate program and, hence, primarily addresses biological EM. However, it is open to students from other disciplines - presently we do not have a full EM course in the physical sciences, although my colleague Dr Thor Bostrom teaches several weeks of EM in various courses for physics, chemistry and engineering undergraduate students.
Apart from this, I will usually run single 2 hour practical sessions for a couple of other undergraduate courses (mainly microbiology, where students look at negatively stained microbes they have isolated from various sources). I'll often be asked to give a half hour (sometimes an hour) introductory EM lecture to these students, prior to the practicals.
I also teach a brief introduction (2-3 weeks) to EM as part of a post-graduate course on cell structure and function in pathology. Other post-graduates needing EM as part of their research are taught individually by staff in our EM Facility, to the level that their projects require.
My full semester EM course is offered in the third year of the B. Applied Science course. This year I have 30 students - the numbers have been increasing over the past few years from a previous average of 20. I usually also have a couple of post-graduate (research) students attending the lectures - these students will be using our EM facilities as part of their research, and I find this is an effective means of giving them some useful background and theory to support their laboratory work.
For this course, I am allocated two by one hour lectures each week (total of 26 hours lectures for the semester). I cover basic instrumentation (TEM, SEM, preparation equipment) and biological sample preparation methods, including cryotechniques. I also include immunolabelling methods, some cytochemistry, microanalysis (Thor Bostrom gives these lectures), a few interesting "non-biological" applications, and a brief introduction to some other imaging methods (confocal, AFM etc).
Practical sessions have been a bit of a "challenge" as student numbers have increased and EM staff numbers have decreased! However, I've now got a system that works, and doesn't cause me (or the equipment) too much stress. I have two by 3 hours practical sessions running each week - ie. 6 hours of my teaching time per week (total of 72 hours of my teaching time for the semester).
Each student attends a total of 24 hours practical sessions for the semester (this fits with our "official" allocation for total practical time for a course of this credit point value). I divide the class into small groups - 3 to 4 students per group, with current student numbers. For preparation lab based pracs (where we have enough space and the students don't need such close supervision), I'll have a number of these small groups attending the same practical session. For EM operation and viewing pracs, I'll only have one small group attending at a time (usually 1 or 1.5 hours of the session for each small group). This has enabled me to run practical sessions almost single-handed - money to employ tutors/demonstrators or other teaching assistants has been a bit difficult to obtain! If anyone would like further information on the practicals I run or on the logistics of this, please contact me by direct email - does anyone else out there run EM practicals for undergraduates??? I haven't seen a rush of emails in response to Robert's questions.
Fortunately, my university is still keen to have students doing "hands-on" practicals in their courses. However, I've certainly had to decrease practical work substantially since I first started teaching in this EM course. A lot of this is due to decreased staff numbers in our EM Facility - a problem that I guess most of us face. I teach other courses (microbiology, parasitology) and encounter the same problems there - it is now very difficult to get competent, experienced people to assist in practical classes, particularly for more advanced or specialised courses.
On the bright side, I've found the students enjoy EM a great deal - and it is one of the few opportunities our undergraduates get to use expensive and (relatively!) modern instrumentation, even if it is only on a very limited scale. I've not had many students who don't show a great deal of enthusiasm when told it is now their turn to operate the TEM or SEM and take some micrographs. This semester, more so than previously, at the end of each practical session quite a number of my students have taken the time to thank me for running the practicals (even after the night classes which finish at 8pm). I figure that has got to be a good sign!!!
Regards Deb ***************************************************** Dr Deborah Stenzel Lecturer (Microbiology) School of Life Sciences and Applications Specialist (Biological) Analytical Electron Microscopy Facility Queensland University of Technology GPO Box 2434 Brisbane 4001 Australia
We are a diagnostic EM lab in surgical pathology, using wet photography processes. If we receive a sample early in the morning, we will have labeled prints in the pathologists hands before 4:30pm the same day. It is basically 8 working hours for most rush cases. If we get the sample later in the day, we will finish the next morning, and prefer to polymerize overnight at 70 deg. C for epon araldite.
In order to speed things up, we put plastic in over at 100 deg C for 1 hr 15 min. The plastic isn't as good, but it does the job. We cure it for 5 minutes.
We offer two EM courses here, one SEM and one TEM. Each is an elective course taken by a mixture of seniors and graduate students, which runs once a year for a full semester. I teach the TEM course, and enrollment is generally anywhere from six to ten students. (More than ten would be a problem in lab.) We have two 50-minute class periods a week, to give the students some foundation in relation to the instrument and electron optics, specimen prep., electron diffraction (we're a materials department), and imaging. There is also a two-and-a-half hour lab session each week, which covers basic operation of the microscope, an introduction to specimen preparation, and indexing of diffraction patterns.
Gill
Dr. Gillian M. Bond Department of Materials & Metallurgical Engineering New Mexico Tech Socorro, NM 87801 (505) 835-5653 gbond-at-nmt.edu
----- Original Message ----- } From: "Robert Fitton" {fittonro-at-luther.edu} To: {Microscopy-at-sparc5.microscopy.com} Sent: Thursday, April 26, 2001 2:40 PM
Hello fellow microscopists:
Does anyone know good protocols for-
1. retina
2. colagen (to prevent shrinkage in skin tissue)
Thanks again, Tim Quinn University of Kansas Museum of Natural History
We have a range of interactive EM courses set out in e-book form. We cover basic SEM, TEM and EDX with more advanced courses in SEM. The pictorial data are in the form of bitmaps and we allow purchasers to use these in their own presentations if they wish.
Full details are on our web site
Steve Chapman Senior Consultant, Protrain For professional training in SEM, TEM and EDX world wide www.emcourses.com
Margaret, I take it from your 'subject' you have a solid block of polymer with a CB dispersion. If this is correct you will have no problem looking at a thin section. Contrast will be low but a low KV ~60 would be good provided you keep the sections thin. For better contrast you can pyrolyze the polymer to improve the contrast. If you would like to do this let me know and I fill you in on the details. Russ Gillmeister Microscopy Xerox Corp. RGillmeister-at-crt.xerox.com
-----Original Message----- } From: Margaret Miller [mailto:MILLERMM-at-uthscsa.edu] Sent: Thursday, April 26, 2001 3:28:PM To: MSA
Does carbon black cause a problem in the TEM? What percautions should I take? Is the use of the cold finger recommended?
Hi! I have a problem with my EDS system. I bought it in 1992 with a SEM. Now, the resolution and the quantification results are getting worse. Maybe it should be enough changing the window and the detector and testing all. I'd like to have more than one offering about repairing manufactory but I don't know to ask about it . Could anyone help me? Thank's in advance.
I also am in a very similar situation, with a detector of about the same age, which appears to have slowly but steadily lost resolution (we're up to nearly 200 eV full width at half max on the Mn K-alpha line).
I have asked the manufacturer how and why this kind of degradation occurs, but haven't gotten a clear answer. I realize there are multiple components of the system from which the problem could be coming (it manifests itself as an approximately constant 50 eV additional fwhm component on all peaks in the spectrum regardless of at which energy). However, I'd like to know if there are reasons why this might be unavoidable in a detector of this age? If not, what are possible causes and how can one prevent it from happening?
Thanks, Wharton
} -----Original Message----- } From: Romina Belli [SMTP:belli-at-science.unitn.it] } Sent: Monday, April 30, 2001 8:04 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: SEM-EDS: change or repair? } } ------------------------------------------------------------------------ } 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! } I have a problem with my EDS system. I bought it in 1992 with a SEM. Now, } the resolution and the quantification results are getting worse. } Maybe it should be enough changing the window and the detector and testing } all. I'd like to have more than one offering about repairing manufactory } but I don't know to ask about it . Could anyone help me? } Thank's in advance. } } Romina Belli
Does anyone have a good protocol for retina fixation for TEM?
I'm dealing with specimens collected in the field and it appears that the initial field fix- "cacodylate and glut" did not preserve the "rods and cones".
A good recomendation for a field fix would be appreciated. Would Karnovsky's work?
I also need to fix frog skin tissue, causing minimal shrinkage. Any suggestions?
I teach three EM classes here which are undergrad and grad level: EM Theory (2 credits Spring Semester), TEM Lab (3 credits Spring semester), and SEM Lab (2 credits fall semester).
The EM Theory class meets 2x 50 mins each week and is lecture only (I bring in lots of hands on show and tells). It covers SEM, TEM, LM, photography (Silver & digital), sample prep, and AEM. I have a number of students who take this without taking the lab courses. The Lab courses require the EM Theory class.
The both labs meet as a group (class limit of 8-9 students) for 2-5 hours on Mondays (Scheduled for 3 hours but fixation days run long). The students then meet one-on-one with a TA for 2hrs on the scope each week. The students “drive” and the TA’s train/assist/watch. Both Labs cover full microscope operation (including alignment, operating parameters, imaging modes, and photography), sample prep (from wet to scope, including embedment, ultramicrotomy, CPD, particulates, staining techniques - 3 separate preps for TEM and 5 for SEM), darkroom, and digital publication plate production. Students take a written exam, an oral exam on the scope (once they “pass” the scope exam they can use the scope without supervision), and turn in a pretty polished portfolio of images. TEM students spend significant amounts of additional time sectioning, staining, and developing film and contact printing.
Yes, these labs are labor intensive (teaching and students), but by the end of either course the goal is that they are qualified TEM or SEM users. Most of the students continue on utilizing the scope for research (most undergrads taking the actually have formal research projects with faculty members that they continue for 1-2 years plus summers, or on to a masters degree). Students are accepted into the lab classes based on research needs.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 352 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu
At Chicago State University I teach a 4 credit hour course in TEM in the fall each year. The students have 2 one-hour lectures per week and 10 hours of supervised lab experience per week. I have usually have 4 to 8 students each year. Since students are scheduled into the lab 2 at a time, this means that someone is with students 30-50 hours per week. For the last few years I have had a technologist assistant who has been able to work with the students, so that we have been able to continue with research and other work during the semester. It really does take two people for this sort of class. In the spring I have taught a 3-credit hour class in SEM. This one has 2 hours of lecture and 6 hours of supervised lab per week. When my students are done with the class, they can prepare specimens competently, use the TEM or SEM to take pictures, so some alignment, understand the principles, and present a simple research project to the department. In the past that was done as a poster or with slides, but in the last few years they have been preparing Power Point presentations.
I teach three EM classes here which are undergrad and grad level: EM Theory (2 credits Spring Semester), TEM Lab (3 credits Spring semester), and SEM Lab (2 credits fall semester).
The EM Theory class meets 2x 50 mins each week and is lecture only (I bring in lots of hands on show and tells). It covers SEM, TEM, LM, photography (Silver & digital), sample prep, and AEM. I have a number of students who take this without taking the lab courses. The Lab courses require the EM Theory class.
The both labs meet as a group (class limit of 8-9 students) for 2-5 hours on Mondays (Scheduled for 3 hours but fixation days run long). The students then meet one-on-one with a TA for 2hrs on the scope each week. The students “drive” and the TA’s train/assist/watch. Both Labs cover full microscope operation (including alignment, operating parameters, imaging modes, and photography), sample prep (from wet to scope, including embedment, ultramicrotomy, CPD, particulates, staining techniques - 3 separate preps for TEM and 5 for SEM), darkroom, and digital publication plate production. Students take a written exam, an oral exam on the scope (once they “pass” the scope exam they can use the scope without supervision), and turn in a pretty polished portfolio of images. TEM students spend significant amounts of additional time sectioning, staining, and developing film and contact printing.
Yes, these labs are labor intensive (teaching and students), but by the end of either course the goal is that they are qualified TEM or SEM users. Most of the students continue on utilizing the scope for research (most undergrads taking the actually have formal research projects with faculty members that they continue for 1-2 years plus summers, or on to a masters degree). Students are accepted into the lab classes based on research needs.
Richard E. Edelmann, Ph.D. Electron Microscopy Facility Supervisor 352 Pearson Hall Miami University, Oxford, OH 45056 Ph: 513.529.5712 Fax: 513.529.4243 E-mail: edelmare-at-muohio.edu
The Microscopy and Imaging Facility of the University of Calgary requires an Electron Microscopy Technician to fill a newly created position within the unit.
The Microscopy and Imaging Facility is a service resource for the University of Calgary supporting users campus wide. The unit works in all areas of science including medicine, biology, chemistry, materials engineering, and geology. The unit is equipped with six electron beam instruments, X-ray micro analyzers, confocal microscope, digital imaging / image analysis and support resources. It is the largest unit of its type in Western Canada.
The successful applicant will have work experience in the operation and routine maintenance of transmission electron microscopes. The demonstrated ability to teach and instruct new users on instrumentation will be a definite asset. Competence in the operation and use of computers and common software packages is a must.
Salary for this position will be dependant on education and experience in the field.
Please send by May 14, 2001 your cv and a covering letter that includes details of your experience that is relevant to this position to: Dr. John Reynolds Department of Cell Biology and Anatomy University of Calgary, 3330 Hospital Dr NW, T2N 1N4, Calgary, Alberta, Canada.
I also am in a very similar situation, with a detector of about the same age, which appears to have slowly but steadily lost resolution (we're up to nearly 200 eV full width at half max on the Mn K-alpha line).
I have asked the manufacturer how and why this kind of degradation occurs, but haven't gotten a clear answer. I realize there are multiple components of the system from which the problem could be coming (it manifests itself as an approximately constant 50 eV additional fwhm component on all peaks in the spectrum regardless of at which energy). However, I'd like to know if there are reasons why this might be unavoidable in a detector of this age? If not, what are possible causes and how can one prevent it from happening?
Wharton
} Hi! } I have a problem with my EDS system. I bought it in 1992 with a SEM. Now, } the resolution and the quantification results are getting worse. } Maybe it should be enough changing the window and the detector and testing } all. I'd like to have more than one offering about repairing manufactory } but I don't know to ask about it . Could anyone help me? } Thank's in advance. } } Romina Belli
Dear Wharton and Romina, Our Kevex detector was installed in the early 80s, and every so often we have had to warm up and pump out the detector to get back the specified resolution (145 eV). Last year, when this didn't work, we sent the detector to Doug Connors (tnas1-at-msn.com), and he cleaned and overhauled it for a good price and got 147 eV resolution. I have no connection to Doug other than as a satisfied customer. Yours,
Bill Tivol Wadsworth Center Albany NY (518) 473-7399 WFT02-at-health.state.ny.us
This is a question about the FEI FIB 200: We are interested in converting our Pt Gas Injection System into Au, but have been told about possible negative effects on the instrument.
Any data/experience/thoughts on Au deposition using FIB (any model) and its effect on the instrument performance will be greatly appreciated!
Thanks! P.S. I realize it is not quite a microscopy question. I have posted it to the FIB-users list, but received only one response so far. Apologize to those who will get this message for the second time.
******************************** Katharine Dovidenko, Ph.D. Scientist UAlbany Institute for Materials and Center for Advanced Thin Film Technology University at Albany SUNY www.albany.edu/cat
251 Fuller Rd. Albany, NY 12203 USA Phone: (518) 437-8781 Fax: (518) 437-8687
} -----Original Message----- } From: Gonzalez-Cabezas, Carlos } Sent: Monday, April 30, 2001 2:37 PM } To: Goheen, Michael P. } Subject: RE: SEM position } } I was asked to post this job opening on the listserver. } } Mike Goheen } } SEM/EPMA tech wanted. The Indiana University School of Dentistry is } looking for a technician for its new digital electron microscopy } laboratory. } A JEOL LV-5310 scanning electron microscope and JEOL 8900R electron probe } microanalyzer are in the process of being installed in a renovated lab in } the IU School of Dentistry. The electron microscopes will have energy- and } wavelength-dispersive spectrometers and are fully digitized. The } laboratory } will serve the research and teaching interests of several units on campus } in } addition to Dentistry including the IU School of Medicine, and the Purdue } School of Science (Departments of Geology, Biology, Chemistry, and } Physics) } and Purdue School of Engineering at Indianapolis. We seek a candidate who } has a range of interests in spatial variations of the microstructure and } composition of materials, and skills in one or more fields such as } computer } technology, electron microscopy, materials science, engineering } technology, } biomedical and geological research. Please contact Dr. Carlos Gonzalez, } Preventive Dentistry Department, IU School of Dentistry. } } Dr. Carlos González-Cabezas, DDS, PhD } Director of the Confocal & Scanning Electron Microscopy Facility } Indiana University School of Dentistry } CGONZALE-at-IUPUI.EDU } } } } -----Original Message----- } From: Goheen, Michael P. } Sent: Wednesday, April 11, 2001 2:17 PM } To: Gonzalez-Cabezas, Carlos } Subject: RE: SEM position } } } -----Original Message----- } From: Gonzalez-Cabezas, Carlos } Sent: Wednesday, April 11, 2001 12:20 PM } To: Goheen, Michael P. } Subject: SEM position } }
{bold} {color} {param} ffff,0000,0000 {/param} {bigger} FYI: This is the last day for early registration for the FRET/FLIM Symposium in San Antonio.=20 Abstracts will be accepted until June 1st.
{/bigger} {/color} {/bold} {bigger} The University of Texas Health Science Center will host a symposium sponsored by Hamamatsu Photonics KK on
{paraindent} {param} left,out {/param} "Spatial resolution of early signalling processes in cells"=20
{/paraindent} Christoph Biskup,=20
{paraindent} {param} left {/param} "Fluorescence lifetime and energy transfer measurements in living cells with a confocal laser scanning microscope and a streak camera"=20
{/paraindent} Robert Clegg,=20
{paraindent} {param} left {/param} "Real-time fluorescence lifetime-resolved imaging - why we do it, how it's done, and applications for biology and medicine."=20
{/paraindent} Michael Edidin
{paraindent} {param} left {/param} "Photobleaching FRET microscopy: practice and theory"
{/paraindent} Enrico Gratton =20
Hans Gerritsen
{paraindent} {param} left {/param} "Fast fluorescence lifetime imaging"=20
{/paraindent} Jesus Gonzalez
{paraindent} {param} left {/param} "FRET Probes and Assays for Drug Discovery"=20
{/paraindent} Brian Herman
{paraindent} {param} left {/param} "FRETing over the apoptotic cascade"
{/paraindent} Thomas Jovin
{paraindent} {param} left {/param} "Extending the capabilities of FRET and FLIM for molecular and cellular biology: phFRET (photochromic FRET), rFLIM (anisotropy FLIM), spectrally-resolved and optical-sectioning FLIM"
{/paraindent} Karsten K=F6nig
{paraindent} {param} left {/param} "Multiphoton microscopy with submicron spatial and picosecond temporal resolution"=20
{/paraindent} Wen-hong Li
{paraindent} {param} left {/param} "Towards the development of highly luminescent lanthanide complexes for FRET and FLIM"=20
{/paraindent} Paloma Mas (substituting for Steve Kay)
{paraindent} {param} left {/param} "Functional interaction of phytocrome B and cryptochrome 2"=20
{/paraindent} Atsushi Miyawaki
{paraindent} {param} left {/param} "Imaging of cellular functions by FREting"
{/paraindent} Ammasi Periasamy
{paraindent} {param} left {/param} "Quantitation of Protein Signals in a Single Living Cell: Wide-field, Confocal, Two-photon and Lifetime FRET Microscopy"=20
{/paraindent} Alexander Sorkin
{paraindent} {param} left {/param} "Interactions of the EGF receptor with adapter proteins during endocytosis" =20
{/paraindent} Roger Tsien=20
{paraindent} {param} left {/param} "FRET based readouts of intracellular messengers and protein interactions"=20
Dear Wharton and Romina, I am running two EDX detectors that are older than Romina's, one 1985 and one of similar age that I bought used. Both still meet their original spec of 149 and 146 eV at Mn Ka. When I have had a degradation of resolution, I turned off the bias, grounded it out with a paper clip on the detector bias connector and warmed up the detector until all the frost was gone from inside the dewar. I used warmed air from a hair drier, blown into a hose down to the bottom, but there other methods that can be used. When the dewar was completely dry, I refillled with liquid nitrogen, let it cool overnight and applied bias the next day. The bias should be on at least one hour before testing the resolution. In one case this cured the resolution, but degraded the holding time for liquid nitrogen, so I then had the dewar re-pumped. I would recommend that step, at least, before buying a new detector. There are also EDX detector repair companies that will diagnose and repair detectors for less than the cost of a new one. I have also had a grounding problem that gave high noise on the detector, because the case on the pre-amp oxidized. A little emery paper cured that. That showed more high dead-time than degraded resolution. At 08:58 AM 4/30/01 -0500, you wrote: } } Dear List, } } I also am in a very similar situation, with a detector of about the same } age, which appears to have slowly but steadily lost resolution (we're up to } nearly 200 eV full width at half max on the Mn K-alpha line). } } I have asked the manufacturer how and why this kind of degradation occurs, } but haven't gotten a clear answer. I realize there are multiple components } of the system from which the problem could be coming (it manifests itself as } an approximately constant 50 eV additional fwhm component on all peaks in } the spectrum regardless of at which energy). However, I'd like to know if } there are reasons why this might be unavoidable in a detector of this age? } If not, what are possible causes and how can one prevent it from happening? } } } Thanks, } Wharton } } } -----Original Message----- } } From: Romina Belli [SMTP:belli-at-science.unitn.it] } } Sent: Monday, April 30, 2001 8:04 AM } } To: Microscopy-at-sparc5.microscopy.com } } Subject: SEM-EDS: change or repair? } } } } ------------------------------------------------------------------------ } } 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! } } I have a problem with my EDS system. I bought it in 1992 with a SEM. Now, } } the resolution and the quantification results are getting worse. } } Maybe it should be enough changing the window and the detector and testing } } all. I'd like to have more than one offering about repairing manufactory } } but I don't know to ask about it . Could anyone help me? } } Thank's in advance. } } } } Romina Belli } 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
Job Posting submitted by Dr. David P. Bazett-Jones
Service Manager, Electron Microscopy Facility
Date Posted: April 30, 2001
Position Status: Full-time, Fixed term
Department: Cell Biology Research Institute
Available: August 1, 2001
Description of the Position: You will share responsibility for the operation and maintenance of transmission and scanning electron microscopes in a new Bioimaging Facility co-sponsored by teaching hospitals in the University of Toronto. The microscopes include an ESEM (FEI/Philips) and a 200 kV TEM (FEI/Philips) equipped with EDX, GIF and cryostage. You will also be responsible for coordination and management
of electron bioimaging services required by investigators of the Hospital for Sick Children Research Institute.
Qualifications: As an ideal candidate, you have completed a M.Sc. in biological sciences, or have completed a B.Sc. with experience in analytical electron microscopy, ultramicrotomy and other sample preparation techniques. Strong computer skills are an asset.
You possess excellent verbal communication and organizational skills. You have the ability to work well independently and in a team.
Hours : 35 hours/week
Salary: $39,848.95 - $50,277.67
Available to: Internal and External Candidates
Deadline: May 9, 2001
Submit Resume to : Erin O’Hare The Hospital for Sick Children 555 University Avenue, Toronto, Ontario M5G1X8
Mananged to get most of the OS updated done this evening. There may be a few hiccups over the next couple of days so be patient. Be prepared for the occasional error message while I reset and fine tune all the system parameters
Cheers.... Nestor Your Friendly Neighborhood SysOp
Dear All Being a botanist and all, I know the cube root of very little about this really, but my understanding is that gold atoms can diffuse into and "poison" semiconductors, and that gold should never be used for specimen coating etc. in any SEM / FIB which may be used to examine or test semiconductors where the semiconductor functionality must be maintained. A) Is this relevant to Katharine's question? B) Is it true or an urban myth?
Chris
----- Original Message ----- } From: "Katharine Dovidenko" {KDovidenko-at-uamail.albany.edu} To: "'Microscopy-at-MSA.Microscopy.Com'" {Microscopy-at-sparc5.microscopy.com} Sent: Monday, April 30, 2001 6:57 PM
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Wharton, It may be as simple as cleaning out your dewar and or outgassing your window. We were losing resolution on our EDAX detector. It was brought up to room temperature for a day or so and the resolution improved greatly. It worth a try if the manufacturer allows it. Clean out any debris from the dewar and make sure your detector isn't powered up during the warm up. Good Luck, Russ Gillmeister Microscopy Xerox Corp. RGillmeister-at-crt.xerox.com
-----Original Message----- } From: Sinkler, Wharton [mailto:WSinkler-at-uop.com] Sent: Monday, April 30, 2001 9:59:AM To: 'Romina Belli'; Microscopy-at-sparc5.microscopy.com
Dear List,
I also am in a very similar situation, with a detector of about the same age, which appears to have slowly but steadily lost resolution (we're up to nearly 200 eV full width at half max on the Mn K-alpha line).
I have asked the manufacturer how and why this kind of degradation occurs, but haven't gotten a clear answer. I realize there are multiple components of the system from which the problem could be coming (it manifests itself as an approximately constant 50 eV additional fwhm component on all peaks in the spectrum regardless of at which energy). However, I'd like to know if there are reasons why this might be unavoidable in a detector of this age? If not, what are possible causes and how can one prevent it from happening?
Thanks, Wharton
} -----Original Message----- } From: Romina Belli [SMTP:belli-at-science.unitn.it] } Sent: Monday, April 30, 2001 8:04 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: SEM-EDS: change or repair? } } ------------------------------------------------------------------------ } 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! } I have a problem with my EDS system. I bought it in 1992 with a SEM. Now, } the resolution and the quantification results are getting worse. } Maybe it should be enough changing the window and the detector and testing } all. I'd like to have more than one offering about repairing manufactory } but I don't know to ask about it . Could anyone help me? } Thank's in advance. } } Romina Belli
We are in the process of updating the Microbeam Analysis Society's membership database. If there are any changes in your personal information (address, phone/fax #'s, email, etc.) as listed in the 2000 directory and you have not made the necessary changes on your 2001 renewal form, please email me with the updated information. Although we will not be printing a new directory this year we would like to keep everyone as current as possible with the membership information.
If you are not a member of MAS and would like to join, please contact me for more information and an application form.
Thanks, Lou Ross MAS Membership Services PMB #141 2101 W. Broadway Columbia, MO 65203-1261 (800) 4MASMEM url: www.microanalysis.org
_______________________________________________________ Send a cool gift with your E-Card http://www.bluemountain.com/giftcenter/
{HTML} {FONT FACE=arial,helvetica} {FONT SIZE=2} We are currently looking for a used ultrcut microtome in good to excellent {BR} condition. {BR} John Hoffpauir {BR} Cooper hospital {BR} Camden NJ {BR} 08107 {BR} 856 757-7781 {/FONT} {/HTML}
Just a general question for the List - are those 10mm x 10mm cylindrical copper (or Al) Jeol type stubs still in wide usage in the SEM community? Do newer Jeol machines still use them? The reason I'm asking is that we have a couple cabinets full of these old stubs from when we used to have a Jeol back in the mid-70's, but of course can't look at them now with our current instrument. I'd like to toss them so we can modify the cabinets to accept our pin-type stubs, and I'd feel better about doing so if I thought it would be hard to find an instrument that could still look at these old ones. Of course, if it turns out that I can't find documentation to indicate what's on these stubs, they'll be going in the bin anyway.
F.C. Thomas MicroAnalysis Facility Geological Survey of Canada (Atlantic) Bedford Institute of Oceanography Dartmouth, Nova Scotia Canada
Workshop Announcement University of California at Santa Barbara
The Department of Molecular, Cellular, and Developmental Biology and the Neuroscience Research Institute are sponsoring an advance course on light microscopy. This 4-day workshop will be offered from August 20 through August 23, 2001 and will consist of lectures and laboratory exercises that will run from 9 am to approximately 5 pm each day. The seminar/workshop will be an intensive lecture/laboratory series that will enable participants to develop theoretical and hands-on expertise with light microscopes. Attendees will closely interact with the instructors while using modern research grade microscopes, cameras, and computers. The seminars and laboratories will cover basic optical theory and how it pertains to increasing contrast (signal to noise ratio) in biological samples. Fundamental techniques such as fluorescence, phase contrast, Nomarski differential interference contrast, and darkfield imaging will be taught and attendees will use microscopes equipped to perform these optical enhancement techniques. In addition, the theory and practice of electronic image acquisition (analog and digital) will be discussed and attendees will work with low-light cameras, digital image processing computers, and morphometric programs. There are five research grade microscopes, five electronic imaging cameras, two computer workstations, and one confocal microscope. With a maximum enrollment of 10 students, there will be ample opportunity to work with all of the microscopes and cameras. For those so interested, intensive hands-on instruction and guidance on the confocal microscope will be provided.
For a fuller description of the workshop please check the web address below. Enrollment forms can be completed online and this workshop provides an opportunity to have a working-vacation in Santa Barbara, California.
The Nikon 990 has a pretty good macro mode and I have found a supplier of a macro lens system that is excellent, but I need a stand that has rough height adjustment like you would find on a stereo scope. Has anyone found a good solution for this presuming that the sample would sit on a table or stand and the Nikon lens would be lowered to the desired height. I know this is typically done with a copy stand, but I find them over-kill (large and provides own illumination). I would use the illumination from my stereo scope so all I need is a small mechanical stage.
Being an electronics technician, I only know enough about optics to be dangerous, but use microscopes often. I am trying to understand what we have, in order to figure out what else could help us out. I would appreciate information for a "layman", or pointers web sites that might enlighten me.
First, I need help understanding the markings on our objective lenses. The question marks indicate what I don't even have an inkling of the meaning.
#1: Zeiss (The maker of our LSM & this lens) Epiplan-NEOFLUAR ( ? ) 100x/0,90 (magnification/numerical aperture) 44 23 80 ( ? )
#3 research devices (The maker) infrared (illumination designed for) Trans 100 IRN ( ? ; mag ; near IR ? ) 0.90 ( NA )
Is it possible to calculate the working distance from the available information?
When a lens is made for infrared use, what is different about it? Are the lens coatings different, or totally absent? Is the lens glass special?
If you read this far, thank you! I thought I would find the answers to these questions, and many more, on an episode of "Soap," but it didn't work. The list is a wonderful pool of knowledge, so don't fail me now!
Chris: if you MUST maintain functionality, it's best not to coat at all. My next choice would be carbon coating, which can be removed by ashing in an O2 plasma. The 3rd choice is Au/Pd, which can be removed with a wet etch of iodine/potassium iodide, but this can attack any exposed aluminum metal on the die.
The biggest problem with using a coating on semiconductors one wishes to remain functional is getting all the coating off to prevent shorts/leakages between the device pins. I don't think it's relevant to Katherine's question as she is concerned about negative effects to her FIB, not the devices.
Most semiconductors have a passivation layer (usually silicon nitride about 1 micron thick) over their surfaces and this protects from unwanted contaminants. I'm no device physicist, but I think the device would have to be heated to a high temperature for any gold implanted in the top atomic layers to diffuse into the active junctions and cause trouble. You want to keep Au out of the fab, but after the passivation is deposited and is intact, the device is fairly impervious to metals sputtered on top. Some older devices used to have Au plated on their backs to help attach them in their packages.
I've used gold (sputter and evaporative) coating over the years to coat semiconductors for SEM exam, and haven't seen any adverse effects. Most of my problems were related to trying to get the coating off afterwards.
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 } -----------------------------------------------------------------------. } } Dear All } Being a botanist and all, I know the cube root of very little about } this really, but my understanding is that gold atoms can diffuse into } and "poison" semiconductors, and that gold should never be used for } specimen coating etc. in any SEM / FIB which may be used to examine or } test semiconductors where the semiconductor functionality must be } maintained. A) Is this relevant to Katharine's question? B) Is it } true or an urban myth? } } Chris } } ----- Original Message ----- } } From: "Katharine Dovidenko" {KDovidenko-at-uamail.albany.edu} } To: "'Microscopy-at-MSA.Microscopy.Com'" } {Microscopy-at-sparc5.microscopy.com} } Sent: Monday, April 30, 2001 6:57 PM } Subject: FIB: Au deposition instead of Pt. } } } -------------------------------------------------------------------- } ---- } } 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 question about the FEI FIB 200: We are interested in } converting } } our Pt Gas Injection System into Au, but have been told about } possible } } negative effects on the instrument. } } } } Any data/experience/thoughts on Au deposition using FIB (any model) } and its } } effect on the instrument performance will be greatly appreciated! } } } } Thanks! } } P.S. I realize it is not quite a microscopy question. I have posted } it to } } the FIB-users list, but received only one response so far. Apologize } to } } those who will get this message for the second time. } } } } ******************************** } } Katharine Dovidenko, Ph.D. } } Scientist } } UAlbany Institute for Materials and Center for Advanced Thin Film } Technology } } University at Albany } } SUNY } } www.albany.edu/cat } } } } 251 Fuller Rd. } } Albany, NY 12203 } } USA } } Phone: (518) 437-8781 } } Fax: (518) 437-8687 } } } }
-- ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Becky Holdford (r-holdford-at-ti.com) 972-598-1291 (pager) DSPS Packaging Development Texas Instruments, Inc. Dallas, TX ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
For macro photography I use a machinist's height gauge. They are usually about 18" high with a smooth, but tight, sliding anvil that can
be machined to accept a camera adapter ring mount. A threaded fine adjustment allows for fine focus. Mounting a Nikon 990 will require that additional weight be added to the gauge's base to prevent the assembly from toppling. Lubricated glass plates and a tiny sandbox can
be used for orienting the sample relative to the camera lens.
Location: Eindhoven, Noord braband, The Netherlands
Question: L.S. I'm doing a survey on bone tissue under the microscope. I want to make the bone tissue visible from a large overview with a regular microscope to a small overview with a (E)SEM. Do you have any experience with this kind of survey's. What kind of problems can I run into (e.g. How can I mark the piece of bone so that I always will see the same spot of bone under the different microscopes?). Do you have reports of other students that have done a study alike this one. At forehand thanks for your time cheers koen kodde student medical engineering
We are doping plastics with high concentrations of dyes and would like to determine the optical transmission properties of these plastics in the wavelength range 350-800nm as a function of thickness of the doped plastic. We do not have the capability to accurately cut thin sections of these plastics to perform these measurements. Ideally we would like to have sections cut at 0.25, 0.5, 0.75, 1.0, 1.5, 2.0, 2.5, 3.0, 3.5, 4.0, 4.5 and 5.0 microns at for a series of dye concentrations in these plastics and have the sections mounted on glass slides. The sections should be free of holes, scratches or other defects. We can cast the plastic to whatever shape is needed but the other dimensions need to be at least 5x5mm.
Is there a service or contract lab out there that can do the sectioning. Interested parties should contact me to discuss further details.
Pleases do not sent to me SU ---------------------- Forwarded by Sumalee Uthaithavorn/BKK/BE/PHILIPS on 2001-05-02 09:43 ---------------------------
milesd-at-US.ibm.com on 2001-05-02 07:33:29 To: Microscopy-at-sparc5.microscopy.com-at-SMTP cc:
Hello All.
Being an electronics technician, I only know enough about optics to be dangerous, but use microscopes often. I am trying to understand what we have, in order to figure out what else could help us out. I would appreciate information for a "layman", or pointers web sites that might enlighten me.
First, I need help understanding the markings on our objective lenses. The question marks indicate what I don't even have an inkling of the meaning.
#1: Zeiss (The maker of our LSM & this lens) Epiplan-NEOFLUAR ( ? ) 100x/0,90 (magnification/numerical aperture) 44 23 80 ( ? )
#3 research devices (The maker) infrared (illumination designed for) Trans 100 IRN ( ? ; mag ; near IR ? ) 0.90 ( NA )
Is it possible to calculate the working distance from the available information?
When a lens is made for infrared use, what is different about it? Are the lens coatings different, or totally absent? Is the lens glass special?
If you read this far, thank you! I thought I would find the answers to these questions, and many more, on an episode of "Soap," but it didn't work. The list is a wonderful pool of knowledge, so don't fail me now!
Greetings to Prof Holland Cheng, Prof Alex Hyatt and all list users.
I worked in a multi-users laboratory and we have facilities equipped for cryo TEM. I have been asked by my colleague to post the following questions:
1. Is it necessary to chemically fixed virus-infected samples for cryo TEM? 2. What are the methods and precautions for cryo TEM of unfixed virus-infected cells? 2. How to dispose the LN2? 3. What are the differences between chemically fixed and unfixed virus-infected samples for cryo TEM?
Hi, I saw your post and thought you might be interested in this...
When you access the Internet, your computer keeps permanent hidden records of your activities! I recently tried EE and I was shocked at what it uncovered on my hard drive.....It actually frightened me. It showed all that I had been doing even though I had deleted it. My advice is to check it out NOW I found it at http://ee1.20m.com Regards, Harry
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