Has anyone out there any ideas on how best to visualise the protofilaments that make up the microtubule subfibres of respiratory cilia? We have tried tannic acid at 1% in the primary fix and at the final dehydration stage.
I would also be interested to hear from anyone with experience of immunogold labelling of the dynein arms.
I apologize for not incuding my address in my last posting. Here is th= e complete e-mail message.
Dear ICEM attendees:
We are looking for sands from around the world to support the GEMS/Proj= ect Micro elementary science program, MICROSCOPIC EXPLORATIONS. If anyone = would like to collect a handful of sand from Cancun for this project it would= be greatly appreciated. If you can send some sand please contact me first= so I can avoid receiving to many samples. The samples can be sent to:
Joe Neilly Abbott Laboratories D-45M, AP31 200 Abbott Park Rd. Abbott Park, IL 60064-3537 voice: (847)-938-5024 fax: (847)-938-5027 e-mail: joe.neilly-at-abbott.com =
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I am trying to prepare virus crystals for HR-SEM. The crystals are 0.1mm or less in size and very delicate. Fixation and dehydration are not a problem. The problems come in the final drying and mounting for viewing.
I have tried drying by gradually replacing the 100% ETOH with Freon 113. However, the crystals float making them very hard to keep track of....also they tend to attach to the walls of the small tube and dry there. I need to get them onto something that can then be put into the SEM.
I normally will have only 2-3 of these crystals so cannot afford to have any lost in the process. They are very hard to see so I hesitate trying to put them onto filter paper...I am afraid they will get lost in the fibers.
I would appreciate any suggestions for ways to handle these guys.
Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
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I have a need to learn how to operate a Biorad confocal microscope -- the scope most available to me is model MRC 1000. I am based in St. Paul, MN, but will travel anywhere for a formal training session. Are such sessions available and, if so, where and when are they? Who do I contact?
Would prefer that you email me directly with your information. Thanks.
We use Epon (EMbed, Polybed, etc) for in situ embedding. Use only ethanol (not propylene oxide) for dehydration. Make sure absolute alcohol is dry by either using fresh bottle or storing it with molecular sieves. (Bake sieves every time bottle is emptied to dry. Let settle after adding ethanol for any sediment to settle out).
Not sure whether Maraglas can be processed this way. Suggestion for determining this to follow.
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
} Hi Folks, } I have been trying to use EMbed-812 for TEM on C.elegans. I am } using an ultracut } E, with a glass knife and I just can not seem to get any thin sections using } this resin.Poly/Bed 812 has worked fine, but is a little too viscous. I have } tried both medium and hard mixtures to no avail. Does anyone have any ideas, } I don't have access to a diamond knife. } } Thank You } } Patrick
Please be more specific about your problem. Are you getting tissue tearout, compression, poor infiltration, or what? Must you use EMbed-812 for some reason?
Caroline Schooley Educational Outreach 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/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/pci.html
} Hi Folks, } I have been trying to use EMbed-812 for TEM on C.elegans. I am } using an ultracut } E, with a glass knife and I just can not seem to get any thin sections using } this resin.Poly/Bed 812 has worked fine, but is a little too viscous. I have } tried both medium and hard mixtures to no avail. Does anyone have any ideas, } I don't have access to a diamond knife. } } Thank You } } Patrick
Please be more specific about your problem. Are you getting tissue tearout, compression, poor infiltration, or what? Must you use EMbed-812 for some reason?
Caroline Schooley Educational Outreach 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/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/pci.html
The New York Society of Experimental Microscopists 1998 Presidential Symposium and The Analytical Imaging Facility of the Albert Einstein College of Medicine
Present
"THE CELLULAR CELL: CREATION AND MAINTENANCE OF AN ORGANIZED CYTOPLASM"
Third Floor Lecture Hall Forchheimer Building, Albert Einstein College of Medicine
September 10, 1998 8:45 AM to 5:30 PM
8:45 AM Registration -- Coffee, 3rd Floor Conference Room
9:15 AM Welcome and Introduction Dr. John Condeelis, President, NYSEM
9:30 AM Dr. Tulle Hazelrigg, Columbia University "Getting the Message to its Destination: Localization of Bicoid mRNA in the Drosophila Oocyte"
10:15 AM Dr. John Condeelis, Albert Einstein College of Medicine "Reciprocal Regulation of mRNA Targeting and Actin Filament Dynamics"
11:00 AM Coffee Break with the Vendors, 3rd Floor Conference Room
11:30 AM Dr. Donald Ingber, Harvard Medical School "Tensegrity: The Mechanical Basis of Cellular Organization"
12:15 PM Dr. Mark Mooseker, Yale University "Myosin Superfamily of Actin Based Motors: Tails of Deafness, Blindness and Seizures"
1:00 PM Lunch at The Analytical Imaging Facility and Vendor Demonstrations, Room F641
3:00 PM Dr. Bruce Schnapp, Harvard Medical School "Biochemical Studies of the RNA Localization Machinery in Xenopus Oocytes"
3:45 PM Dr. Robert Singer, Albert Einstein College of Medicine "Molecular Biology through the Microscope: Intracellular Travels of an RNA"
4:30 PM Conclusion and Perspective Dr. Robert Singer
4:45 PM Open House wine and cheese reception at the Analytical Imaging Facility, Room F641
For additional information see: http://www.ca.aecom.yu.edu/aif/directions.htm
**************************************************************************** Frank Macaluso tel: 718-430-3547 Analytical Imaging Facility fax: 718-430-8996 Albert Einstein College of Medicine e-mail: macaluso-at-aecom.yu.edu 1300 Morris Park Avenue Bronx, NY 10461 ****************************************************************************
MMMS will host a meeting on Friday, October 9, 1998 at Purdue Universit= y in Lafayette, Indiana. The focus of the meeting will be Materials Scienc= e. Details will follow, but mark your calendars now!
Jane A. Fagerland, Ph.D. Dept. of Microscopy and Microanalysis Abbott Laboratories Abbott Park IL 60064 (847) 935-0104 =
I am double labelling plant ovules with a view to observing nuclei and cytoskeleton within the cells of the embryo sac. I am using fixed tissue embedded in a low melting point wax (Steedmans wax) which is removed with alcohol prior to double labelling for tubulin (antibodies - FITC tag) and DNA (Hoechst).
Anti-fade agents (glycerol- PDA) and Citifluor work quite well for FITC but not at all for Hoechst, which fades and/or becomes non-specific throughout the tissue within 24 hours. A 'no-antifade' solution which is made up in glycerol (a mixture of biocarb buffers) and adjusted to pH 8.6 seems the best so far, but this is still not adequate to collect images with uv after 3-4 days.
I'd be grateful for any help
Meredith Wallwork (Dr) Department of Horticulture, Viticulture and Oenology Waite Campus University of Adelaide Sth Aust
Delicate specimen that defy handling during CPD and similar methods often are well preserved by a solvent drying method:
Line a glass Petrie dish with a double-layer of filter paper. Saturate the paper with chloroform (somebody ought to try how other solvents perform). Place a microscope slide onto the filter paper to keep the specimen off the paper. Sit the dehydrated, wet (absolute ethanol) specimen, which may be on a coverslip, a piece of mica etc onto the slide. Cover the Petrie dish and refrigerate for two days. Don't open Petrie dish until it has warmed to at least room temperature (incubator?) Proceed with sputter coating . . .
The method relies on very slow removal of the solvent saturated atmosphere within the dish and this lengthy time allows most of the remaining, trapped water molecules to leave the specimen without high pressures that distorts and shrinks specimens. I have used this method with microscopic nematodes; its not always perfect but for these critters its about the best means available. Certainly, its easy and avoids losing specimens. Cheers Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au *****
On Tuesday, 1 September 1998 23:52, Debby Sherman [SMTP:sherman-at-btny.purdue.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.ht } ml } ---------------------------------------------------------- } -------------. } } } I am trying to prepare virus crystals for HR-SEM. The } crystals are } 0.1mm or less in size and very delicate. Fixation and } dehydration are not } a problem. The problems come in the final drying and } mounting for } viewing. } } I have tried drying by gradually replacing the 100% } ETOH with Freon } 113. However, the crystals float making them very hard to } keep track } of....also they tend to attach to the walls of the small } tube and dry there. I } need to get them onto something that can then be put into } the SEM. } } I normally will have only 2-3 of these crystals so } cannot afford to } have any lost in the process. They are very hard to see } so I hesitate } trying to put them onto filter paper...I am afraid they } will get lost in the } fibers. } } I would appreciate any suggestions for ways to handle } these guys. } } Debby Sherman, Manager Phone: 765-494-6666 } Microscopy Center in Agriculture FAX: 765-494-5896 } Dept. of Botany & Plant Pathology E-mail: } sherman-at-btny.purdue.edu } Purdue University } 1057 Whistler Building } West Lafayette, IN 47907-1057 }
Serge Oktyabrsky raised an interesting question about tapes suitable for cleaving HOPG (highly ordered pyrolytic graphite) to provide single crystal graphite support films for TEM. So far as I'm aware, there isn't one. Certainly the published references for this technique which I've seen use solvents which pose a considerable risk to laboratory workers and their neighbors.
If somebody comes up with an adhesive which adequately sticks to HOPG to cleave it well and then leaves no residue, I'd appreciate knowing about it.
Disclaimer: SPI Supplies sells HOPG, and we have an obvious interest in promoting its use. Furthermore, if we had a technique for reliable production of single crystal carbon support films from cleaved HOPG, we'd add them to our list of products.
Andy
Andrew W. Blackwood, Ph.D. Structure Probe, Inc. P.O. Box 656 West Chester, PA 19381-0656 Ph: 1 610 436 5400 FAX: 1 610 436 5755 e-mail: ablackwood-at-2spi.com WWW: http://www.2spi.com
Have some sponge in here to thin section and eventually IEM. Any suggestions on getting rid of the spicules. They are of the silica variety and HF is not my first choice. Thanks
} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} {} { GO GATORS Scott D. Whittaker 218 Carr Hall EM Technician Gainesville, FL 32610 University Of Florida ph 352-392-1184 ICBR EM Core Lab fax 352-846-0251 sdw-at-biotech.ufl.edu http://www.biotech.ufl.edu/~emcl/ The home of " Tips & Tricks "
Regarding a recent discussion about physical characteristics of hair: I just recieved a notice of a CRC book 'Atlas of Human Hair. Microscopic Characteristics', by Robert R. Ogle, Jr., and Michelle J. Fox. The book is targeted to forensic researchers and practitioners.
The book is due out Feb. 1999, and is priced at $99.95. ISBN: 0-8493-8134-7
I have deleted the original message, so cannot send this to the young student's contact person. If you would like the complete description of the book, contact me and I can fax (or mail) the ad to you.
Maureen Petersen
************************************************************************ Maureen Petersen Department of Plant Pathology 1453 Fifield Hall University of Florida
If you have access only to glass knives, you should immediately consider a mixed resin embedding. The "812s" used with DDSA and NMA are too hard on glass edges. Even with the best glass knife, relatively few thin sections can be cut. Not so with the mixed resin embedments. Once I spent 18 months sectioning tough muscle tissue with glass knives, and was able to get multitudinous sections before the glass edge wore out. It has been suggested to me (but I have not proof of this) that the molecular structure of NMA wears the glass edge too fast. Below is the formulation for this mixed resin which we use today frequently.
Polymerize 48 hours at 60C. Test your block. For a harder block, heat it at 95C for an hour, or put it back at 60C for another 24 hours. You may vary the cutting consistency of your block by adjusting (slightly) the dibutyl use 1/2% or 1%. I have used 2%, but found it too soft. It is a matter of preference and also your final decisions on what sort of sections you need - Silver? Gold? If you go to mixed resin embedding, be sure to mix the resin monomers very well. Keep them mixed. Do not let them just "sit". Keep your tissues on a rotator. Lenghten your embedding times. You are now dealing with the viscous Araldite. Use acteone as an intermediary and do several changes, making sure that all the acetone is out of the tissue. My phone # is 303-871-3026. Bye, Hildy
P.S. DMP-30 has gotten a lot of bad press lately. We have not run tests as to the substitution ratios for the better BDMA. MSA members cannot agree on the quantities either. quantities.
How about trying Nucleopore filters to trap your paticles. These filters are very smooth and I have used them successfully to entrap samples for SEM. There are are a variety of pore sizes available. I know they are available from SPI Supplies--probably other suppliers carry them also.
} Date: 01 Sep 98 08:51:42 -0500 } From: Debby Sherman {sherman-at-btny.purdue.edu} } Subject: virus crystals for SEM } To: "message to: MSA list" {microscopy-at-sparc5.microscopy.com} } X-Mailer: QuickMail Pro 1.5.3 (Mac) } Reply-To: Debby Sherman {sherman-at-btny.purdue.edu} } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
We have found that the SlowFade Light Antifade Kit from Molecular Probes does not quench shorter wavelength fluorophores such as DAPI and Hoechst. Their Prolong Kit has also produced superior results in some applications such as unfixed GFP expressing tissue. See their helpful handbook on line at: http://www.probes.com/handbook/toc.html go to Chapter 26.1 http://www.probes.com/handbook/ch26-1.html#ProLong
--just a happy customer
} Date: Wed, 2 Sep 1998 16:49:21 +0930 (CST) } From: Meredith Wallwork {mwallwor-at-waite.adelaide.edu.au} } Sender: Meredith Wallwork {mwallwor-at-waite.adelaide.edu.au} } Reply-To: Meredith Wallwork {mwallwor-at-waite.adelaide.edu.au} } Subject: re: antifade agents } To: Microscopy-at-sparc5.microscopy.com } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Perhaps the following correlation would be useful to the student: hair straightness and cross section are related. Straight hair is round in cross section, curly hair is more elliptical, and tightly curled hair is flatter still.
Leonard Corwin Fort Dodge Animal Health Princeton NJ
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Regarding a recent discussion about physical characteristics of hair: I just recieved a notice of a CRC book 'Atlas of Human Hair. Microscopic Characteristics', by Robert R. Ogle, Jr., and Michelle J. Fox. The book is targeted to forensic researchers and practitioners.
The book is due out Feb. 1999, and is priced at $99.95. ISBN: 0-8493-8134-7
I have deleted the original message, so cannot send this to the young student's contact person. If you would like the complete description of the book, contact me and I can fax (or mail) the ad to you.
Maureen Petersen
************************************************************************ Maureen Petersen Department of Plant Pathology 1453 Fifield Hall University of Florida
by heinlein.acpub.duke.edu (8.8.5/Duke-4.6.0) with ESMTP id PAA05118; Wed, 2 Sep 1998 15:50:52 -0400 (EDT) Received: (from saram-at-localhost) by soc11.acpub.duke.edu (8.8.5/Duke-4.4) id PAA05896; Wed, 2 Sep 1998 15:50:51 -0400 (EDT)
On Wed, 10 Jun 1998, Ellis, Sarah wrote:
} Date: Wed, 10 Jun 1998 13:45:39 +1000 } From: Ellis, Sarah {s.ellis-at-pmci.unimelb.edu.au} } To: Microscopy Listserver {Microscopy-at-sparc5.microscopy.com} , } Microscopy-at-sparc5.microscopy.com } Subject: Fixing and processing cell colonies grown between agar sheets. } } ------------------------------------------------------------------------ } 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! } } Can you help us? A student is growing very small colonies of cells and } she wishes to view their ultrastructure. There are about 50 cells per } colony and about 4 colonies per 3cm polystyrene petri dish. Our problem } is that the colonies are growing between two layers of agar. The bottom } layer is 0.5% agar in PBS and the top layer is 0.33% agar in PBS. The } colonies break up and float away during processing and the agar just } moves around. The colonies are not attached/embedded in the agar. } We have tried (1)cutting around the colonies and sucking the whole lot } up (agar + colony) and treating it as a pellet but the cells are } impossible to find in the resin, and (2)we have tried to just gently } fix the mass and process it as a whole but we end up with no cells as } the colony breaks up and the cells disperse. } Is there anyone out there who could offer a suggestion. } Thanks } } Sarah Ellis } } } Research Division } Peter MacCallum Cancer Institute } Locked Bag #1 } A'Beckett Street } Melbourne, Victoria 3000 } Australia } } Phone 61-3-9656 1244 } Fax 61-3-96561411 } Email s.ellis-at-pmci.unimelb.edu.au {mailto:s.ellis-at-pmci.unimelb.edu.au} } I suggest:
Keep agar layers as thin as possible (barely covering the plate and then the inoculated cells. It will be only a couple mm thick.
Infiltrate/embed the whole agar layer in situ as you would for an adherrent culture.
Use a microscope to locate the cells, circling the colony with a thin magic marker on the bottom of the plate.
After baking and before you peel up the agar/resin layer, transfer the circle to the upper surface of the layer. You can even put this circle of resin back under a microscope to see the cells which will be slightly brown from the osmium.
THEN cut out the colony and glue onto a blank stub.
We have done this successfully with very small colonies.
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
We trialled a number of antifade solotions, both commercial and DIY last year . Far and away the best for our purposes was Molecular Probes Prolong, Also the most expensive, but you get what you pay for, I suggest you give it a go. Pete Smith AgResearch Wallaceville Upper Hutt New Zealand
I read some of the other responses to your problem and I have used the Prolong Anti-Fade from Molecular Probes and love it. However, we did come up with a very cheap media that has worked very well for double and triple labels in cojuction with a DAPI stain.
70% glycerol 25% .5M tris pH 9.0 5% N-propyl gallate
Heat in a boiling water bath to dissolve the n-propyl gallate. Cool and pH to 7.4 Keep light tight at 4 degrees C. Use as little as neccessary to cover the coverslip.
Bob Derm Imaging Center U of W Seattle
On Wed, 2 Sep 1998, Meredith Wallwork 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 double labelling plant ovules with a view to observing nuclei and } cytoskeleton within the cells of the embryo sac. I am using fixed tissue } embedded in a low melting point wax (Steedmans wax) which is removed with } alcohol prior to double labelling for tubulin (antibodies - FITC tag) and } DNA (Hoechst). } } Anti-fade agents (glycerol- PDA) and Citifluor work quite well for FITC } but not at all for Hoechst, which fades and/or becomes non-specific } throughout the tissue within 24 hours. A 'no-antifade' solution which } is made up in glycerol (a mixture of biocarb buffers) and adjusted to pH } 8.6 seems the best so far, but this is still not adequate to collect } images with uv after 3-4 days. } } I'd be grateful for any help } } Meredith Wallwork (Dr) } Department of Horticulture, Viticulture and Oenology } Waite Campus } University of Adelaide } Sth Aust } } }
When I was in second year, a long time ago I did a Scanning Tunneling Microscopy (STM) exoeriment as part of my materials lab techniques. One of the samples that we studied was HOPG, and we used normal sticky tape to cleave a new surface. The technique was to press a piece of tape onto the HOPG and then just rip it off.
Perhaps you could remove the film on the tape by dissolving the tape adhesive in ethanol, float off the graphite film and then dry it by baking gently. If the gods are smiling, hopefully it will be a single crystal graphite support film.
Don't know if it helps but thats my 2 cents worth!
I have a client that I am assisting in finding a used SEM for a new metallurgical lab. We have been looking at a couple, but I thought there might be some citizens of this list that may be looking to dispose of some older equipment for a good price. LaB6 would be nice but not necessary. EDS also desired, preferably thin window.
Please respond by email if you have an instrument you'd like to sell. Allen R. Sampson Advanced Research Systems 317 North 4th. Street St. Charles, IL 60174 PH 630.513.7093 FAX 630.513.7092 Email: ars-at-mcs.net WWW: http://www.mcs.net/~ars Analytical instrument maintenance services
Stowe and Robinson report about reducing beam scattering in conventional Low Vacuum SEM's (Scanning, Vol. 20, 57-60). Are there any experiences using Helium in an ESEM from Electroscan or Philips with a special ESEM-detector? Is the ionization efficiency high enough to get a good performance for amplifying the electrons coming from the sample? How is the image quality compared to e.g. water vapor?
Kind regards
Rainer Ziel ------------------------------------------------------------- Dipl.-Phys. Rainer Ziel Akzo Nobel Central Research ACR-O/RMG-EM D-63784 Obernburg Germany
As I remember from about five years ago, a student with whom I shared a lab was cleaving HOPG for a STM resolution test specimen by using some form of sticky tape. Dr Mark Aindow was his (and my) PhD supervisor and he can be reached on m.aindow-at-bham.ac.uk, perhaps he would remember the details as he oversees the STM/AFM facility. I was under the impression, however, that this was not some new technique invented by them but something that had been done previously by other STM researchers.
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ Ian MacLaren, Tel: (44) (0) 121 414 3447 IRC in Materials for FAX: (44) (0) 121 414 3441 High Performance Applications, email: I.MacLaren-at-bham.ac.uk The University of Birmingham, http://web.bham.ac.uk/I.MacLaren/ Birmingham B15 2TT, England. ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
I work with Atriplex nummularia and I want to examine epidermal cell patterns. This specie has a proffusion of glandular hairs in both surface= s. I read in a paper something about use cellulose acetate film with acetone to make impressions of the leaf.=20
Someone know something different and simplest about this subject? I also need to know what to do to take off these trichomes. Thank in advance.
Rejane
Rejane Magalh=E3es Pimentel Galindo =20 ggalindo-at-elogica.com.br Universidade Federal Rural de Pernambuco Av. Boa Viagem, 6592/602 FAX: 55 (081) 4416177 51130-000, Recife, Pernambuco, Brasil
Sory, I forgot the final of this message. Here it is complete.
I work with Atriplex nummularia and I want to examine epidermal cell } patterns. This specie has a proffusion of glandular hairs in both surfaces. } I read in a paper something about use cellulose acetate film with aceto= ne } to make impressions of the leaf.=20 } =20 } Someone know something different and simplest about this subject? } I also need to know what to do to take off these trichomes.
Aside this, I want to obtain the vein impressions; I need a simple method to highlight the leaf venation. } Thank in advance. } =20 } Rejane
Rejane Magalh=E3es Pimentel Galindo =20 ggalindo-at-elogica.com.br Universidade Federal Rural de Pernambuco Av. Boa Viagem, 6592/602 FAX: 55 (081) 4416177 51130-000, Recife, Pernambuco, Brasil
Dear listers, we need to do service on JEM 2010 that I suppose is normaly done by = service people as there is nothing about it in the manual. Namely that = is dismounting and clearning of sample lock. Due to some reasons we are = not able to call for JEOL ingineer and have to do all by ourselfs. Does = anyone know of service (persons) whom we could ask for advice? Schemes = and drawings would help greatly.
Before we begin a thread on how to cleave HOPG, may we all please recall that the question was how to cleave it without leaving any tape residue. To cleave using almost any tape is a matter of technique. To cleave without leaving any tape residue is the problem for which the original writer (Serge Oktyabrsky) is seeking a solution.
Andy
Andrew W. Blackwood, Ph.D. Structure Probe, Inc. P.O. Box 656 West Chester, PA 19381-0656 Ph: 1 610 436 5400 FAX: 1 610 436 5755 e-mail: ablackwood-at-2spi.com WWW: http://www.2spi.com
Here is how we were able to get good 'screen shoot' color slides:
Film: Kodak Ektachrome color slide film; ASA 100
Settings: Camera -- Nikon with tripod and cable shutter release; f/2.8 (wide open); 1/2 and 1 sec. No flters.
Computer screen settings: At the 1 sec. exposure, dimming the brightness/contrast on the screen slightly, gave good results. Recommend shooting at 1/2 sec. with a screen appearance good 'to the eye' and 1 sec. with slight screen dimming. Take these two shots on each subject for a choice (often both slides are usable).
Room conditions: Total darkness -- no room lights; close door. This eliminates any glare on computer screen by room light.
Hope this is helpful. Please ask about anything I may have overlooked. Gerald Harrison ======================================================================== At 04:29 AM 9/3/98 -0700, you wrote:
} } I would like to photograph my multi-sync computer monitor screen using } 35mm color slide film. } } Assuming it is worth a try... does anyone have suggestions regarding: } 1)film type 2)shutter speed 3)filters? } } Thank you. } } Bart Cannon } Cannon Microprobe } }
Greetings. Does anyone know how long you can keep Formvar in solution? I need to make perfect films. I have a lot of Formvar in Ethylene Dichloride, but it has been on the shelf unopened since 1990. Can I use it?
} } } I would like to photograph my multi-sync computer monitor screen using } 35mm color slide film. } } Assuming it is worth a try... does anyone have suggestions regarding: } 1)film type 2)shutter speed 3)filters? } } ...
The hardest part may be capturing the dynamic range of the monitor ... you may want to reduce its contrast a bit ... but its difficult to know how much.
Depending on your subject matter ... you may also want to create a standard gray image to meter on ... gray level (R,G,B) = 70 ought to be pretty close. An equivelent and/or comparison can be made by metering "white" with an ASA setting = 1/5 of the film speed (... a copy stand trick ...)
The lens should be a portrait type ... e.g., 100-135mm
The shutter speed should capture many screen refreshes ... e.g., 1/8 sec ... and turn the room lights off.
... hope this helps :o)
cheerios, shAf
{} /\ {\/} /\ {\/} /\ {\/} /\ cogito, ergo zZOooOM /\ {\/} /\ {\/} /\ {\/} /\ {} Michael Shaffer, R.A. - ICQ 210524 Geological Science's Electron Probe Facility - University of Oregon mshaf-at-darkwing.uoregon.edu - http://darkwing.uoregon.edu/~mshaf/
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Debby,
How about cryo-SEM technique? The virus crystals in suspension are absorbed on substrata, fast frozen, partial freeze-dried, cryo-coated with a thin layer of metal, then viewed in a cryo-SEM. I have used this protocol for many years to look at individual viral particle by our high-resolution cryo-SEM.
Ya Chen
Ya Chen
======================================================================== \ / Integrated Microscopy Resource (IMR)-- \ / __ a NIH Biomedical Research Resource TEL: 608-263-8481 \/ / / University of Wisconsin-Madison FAX: 608-265-4076 / / / 1675 Observatory Drive #159 / /__/_ Madison, WI 53706 Email: ychen14-at-facstaff.wisc.edu ======================================================================== IMR WWW Home Page: http://www.bocklabs.wisc.edu/imr/home.htm
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RE} Anybody knows about the_ 9/3/98 10:18 AM Dear Gehoon-at-plaza1....,
I am not sure what your real question is. However, you may want to be awa= re that phase contrast is major contrast technique used in light microsco= py. The technique was developed for telescopes by Frits Zernike (1934) an= d later applied to the microscope by Kohler and Loos at Zeiss.
Phase contrast provides an efficient method for enhancing the contrast of= specimens whose refractive indices are very similar and/ or transparent = (or nearly so). The technique does not apprciable lessen the resolution o= f the optics. The technique is especially used on living cells.
I hope this abbreviated answer will serve you. You may want to visit the = library and pick up any modern light microscopy textbook in the later hal= f of the 20th Century for a more detailed explaination.
john shane
-------------------------------------- the internet for 5 hours, and still don't have the answer. If can anybo= dy help me, please e-mail me at gehoon-at-plaza1.snu.ac.kr =0A {!DOCTYPE HTML PUBLIC "-//W3C//DTD W3 HTML//EN"} =0A {HTML} =0A {HEAD} =0A =0A {META content=3Dtext/html;charset=3Dks_c_5601-1987 http-equiv=3DConten= t-Type} =0A {META content=3D'"MSHTML 5.00.0518.7"' name=3DGENERATOR} =0A {/HEAD} =0A {BODY bgColor=3D#ffffff} =0A {DIV} {FONT color=3D#000000 face=3D size=3D2} I am a freshman studying d= entistry in=20 =0ASeoul, Korea. Yesterday my professor gave us a question which we= were=20 =0Asupposed to answer via e-mail as quick as we could. The problem = was=20 =0A"Why is there the 'phase contrast' in PCM?" I looked u= p all the=20 =0Areferences available to me, and searched all over the internet for 5 h= ours, and=20 =0Astill don't have the answer. If can anybody help me, please e-ma= il me at=20 =0A {A=20 =0Ahref=3D"mailto:gehoon-at-plaza1.snu.ac.kr"} gehoon-at-plaza1.snu.ac.kr {/A} {/F= ONT} {/DIV} =0A {/BODY} {/HTML} =0A
by bubba.NMSU.Edu (8.9.0/8.9.0) with ESMTP id JAA17863; Thu, 3 Sep 1998 09:28:10 -0600 (MDT) Received: from nestor.NMSU.Edu (nestor.NMSU.Edu [128.123.34.146]) by dns1.NMSU.Edu (8.9.0/8.9.0) with ESMTP id JAA11839; Thu, 3 Sep 1998 09:27:04 -0600 (MDT) Received: from microscope.nmsu.edu (microscope.NMSU.Edu [128.123.5.85]) by nestor.NMSU.Edu (8.8.6/8.7) with SMTP id JAA139504; Thu, 3 Sep 1998 09:27:05 -0600 Message-Id: {3.0.6.32.19980903092600.007aa710-at-cnmailsvr.nmsu.edu} X-Sender: rtindell-at-cnmailsvr.nmsu.edu X-Mailer: QUALCOMM Windows Eudora Light Version 3.0.6 (32)
At 04:29 AM 9/3/98 -0700, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Bart,
Although it's bean a while since I've done this, I believe that most computer screens will photograph fairly well on regular daylight slide film (somebody please correct me if I'm wrong), which gives you a pretty wide choice. The slower speed films give you finer grain. A good bet might be one of the 100 speed Fujichromes or Ektachromes.
As to shutter speed, you will want to use a shutter speed slower than the screen refresh rate to avoid bands. You'd be safe with 1/30 of a second or slower for almost any screen, including your home tv.
Good luck.
Randy
} Randy Tindall Electron Microscope Laboratory Box 3EML--Biology New Mexico State University Las Cruces, NM 88003
Transmission microscopy in some contexts might be thought of as the taking of transmitted wave pictures of foils at high magnification -- a kind of seriously enlarging xerox machine that only works for "transparencies". Some microscopes in the past, I believe, may have been designed with this perspective in mind.
The kinds of electron scattering visitation, specimen modification, and analysis, done especially with crystalline specimens, is of a different sort. It rather seems that the microscope is simply a kind of "suit" that you can put on, and that allows one to visit places as a nano-human, and to perform various kinds of electron scattering and data recording activities. Different people, when they visit the tiny places accessible with such "suits", do very different things when they get there, and bring back different stories, much as visitors to the same south sea island might go to the same place but have different experiences as well. Moreover, unlike xerox machines, a suit is merely an extension of a human, and although it may have some personality, it does little by itself.
In this latter spirit, I offer the alternative meaning for the acronym "TEM" given in the subject to this message. For a bit further development of this metaphor, you might enjoy the announcement by this same name linked to our TEM course page* this semester.
As long as you keep it in the dark, it can last a long time. When exposed to light, you tend to form HCl molecules in the dichlorethane weakening the formvar.
Henk
At 10:46 AM 9/3/98 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I can't give you the exact answers you are looking for, but hopefully the following can point you in the right direction. At the '99 MSA meeting in Atlanta, there was a session on ESEM. Several papers were given by the group from the Cavendish Laboratory, especially one on SEM at freezer temperatures by A.L. Fletcher, ... & A.M. McDonald. In an attempt to maintain the proper humidity without icing or thawing, this group used other gases, and determined that N2 was minimally acceptable in terms of obtaining desired imaging SEs. Discussion following the paper included comments on the scattering and SE generation that could be expected from lower atomic number gases, including He. I would suggest your contacting this group. Additionally, I would recommend that you do a literature search for the work of K. Rudiger-Peters and the various publications he has on the physical characteristics and SE generation in the ESEM. I think most of those papers were in the journal Scanning. The principal author on the ESEM is of course Danilatos, and his publications span from 1982 to 1990. Since I do not have ready access to those publications (everything is in boxes, no longer arranged as per my reference manager database) I can't give you figures or exact references even, but I believe that he did publish on different gases. Hope this helps.
Roger Moretz Toxicology Boehringer Ingelheim Pharmaceuticals, Inc.
} -----Original Message----- } From: Ziel, R. (Rainer) [SMTP:Rainer.Ziel-at-akzonobel.com] } Sent: Thursday, September 03, 1998 2:43 AM } To: 'MSA' } Subject: ESEM: Use of Helium Gas } } ---------------------------------------------------------------------- } -- } 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 } ---------------------------------------------------------------------- } -. } } } Stowe and Robinson report about reducing beam scattering in } conventional } Low Vacuum SEM's (Scanning, Vol. 20, 57-60). Are there any experiences } using Helium in an ESEM from Electroscan or Philips with a special } ESEM-detector? Is the ionization efficiency high enough to get a good } performance for amplifying the electrons coming from the sample? How } is } the image quality compared to e.g. water vapor? } } Kind regards } } Rainer Ziel } ------------------------------------------------------------- } Dipl.-Phys. Rainer Ziel } Akzo Nobel Central Research } ACR-O/RMG-EM } D-63784 Obernburg } Germany } } Tel: (06022) 81-2645 } Fax: (06022) 81-2896 } E-mail: Rainer.Ziel-at-AkzoNobel.com }
SE, You can also make bead slides, bought from molecular probes in many sizes and emissions. A little poly-l-lysine to make them stick to the coverslip and you are good to go.
At 09:26 AM 9/3/98 -0600, Tindall wrote: } Although it's bean a while since I've done this, I believe that most } computer screens will photograph fairly well on regular daylight slide film } (somebody please correct me if I'm wrong), which gives you a pretty wide } choice. The slower speed films give you finer grain. A good bet might be } one of the 100 speed Fujichromes or Ektachromes. } } As to shutter speed, you will want to use a shutter speed slower than the } screen refresh rate to avoid bands. You'd be safe with 1/30 of a second or } slower for almost any screen, including your home tv.
I don't think 1/30th will be long enough, especially for TV's. They scan at 60 frames per second but only refresh half the lines per pass, so it takes 1/30th of a second to get one whole image. Now if the camera shutter is exactly 1/30th second, then you would get one and only one complete pass.
We tried a similar exercise on one of our EDS monitors years ago and used too fast a speed. I think I figured we captured 4 and some scans. We could see a brighter band on the slide where 5 frames had been scanned compared to where only 4 frames had been scanned.
So longer is better, and you will probably want to use a tripod for making sure you have the same setup between shots anyway.
I used my 60mm macro (Nikon calls macro lenses micros), and just trusted my light meter in the camera, and used it on program mode, for perfect results with the Nikon F4 -35mm camera. With the macro lens, you can get close enough to fill the frame with the image of the computer screen. You just have to be careful to turn off the room lights to avoid glare, and have the camera parallel with the screen. No filters are necessary, and you can use ordinary daylight slide film.
} ---------- } From: shAf[SMTP:mshaf-at-darkwing.uoregon.edu] } Sent: 3 September, 1998 10:08 } To: cannonmp-at-accessone.com; MSA Listserver } Subject: RE: Photos of Comptr Scrn } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
We wish to buy used AFM systems and components made by Digital Instruments:
NanoScope II NanoScope III Contact mode or Multimode AFM or Dimension AFM AFM base AFM scanners AFM optical head Accessories and tools used with NanoScope equipment, including fiberoptic illuminator, stand for monocular microscope, vibration isolation pad, acoustic shroud.
Example: If you originally had a NanoScope II AFM and upgraded to a NanoScope III AFM to do TappingMode, you may still have a contact mode AFM base and optical head that you are not using.
If you have equipment to sell, or if you know someone who may have such equipment, please contact me offline (directly) at the address shown below. Do not respond to this discussion group.
Thank you.
Don Chernoff
Advanced Surface Microscopy, Inc. E-Mail: asm-at-indy.net 6009 KNYGHTON RD. Voice: 317-251-1364 INDIANAPOLIS IN 46220 Toll free: 800-374-8557 (in USA) web: http://www.a1.com/asm Fax: 317-254-8690 (note: "a1"= letter "a", numeral "1")
Dear Bart, } } I would like to photograph my multi-sync computer monitor screen using } 35mm color slide film. } } Assuming it is worth a try... does anyone have suggestions regarding: } 1)film type 2)shutter speed 3)filters? } I have been successful at photographing monitors; however, there are distortions due to the difference between the camera-screen distances to the center or edge of the screen. Bearing this in mind, one would like to use a high f-number for a large depth-of-field. I used (1) ektachrome 100 professional film, (2) the auto-exposure setting (an exposure much longer than the refresh time is good so that many frames contribute to the image), and (3) no filter. If distortions are not acceptable, try a tele- photo lens, but be prepared for a very long exposure. Good luck. Yours, Bill Tivol
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Greetings. } Does anyone know how long you can keep Formvar in solution? I need to make } perfect films. I have a lot of Formvar in Ethylene Dichloride, but it has } been on the shelf unopened since 1990. Can I use it? } } Sally Shrom } } } Hi! Formvar solutions deterioate due to atmospheric moisture and age. This causes holes. If not holes, then "measles" (light areas in the film). It is not worth ruining sections because one has used old Formvar solution. Buy or make new from powder. Make a few films and check in the TEM for stability, thickness, etc., before making a thousand. Bye, Hildy
Sally Shrom wrote: } } } Greetings. } Does anyone know how long you can keep Formvar in solution? I need to make } perfect films. I have a lot of Formvar in Ethylene Dichloride, but it has } been on the shelf unopened since 1990. Can I use it? } } Sally Shrom Hi Sally,
I use Formvar in Cloroform solution in a well stopped glass ( 100 ml, 0.25%). I am using the solution at 1 year and obtained excellent films. With a pen, mark the level of the solution and if it evapore, refill with Cloroform. The Butvar give excellent films too and it easy to make very fine films. Rinaldo Pires dos Santos UFRGS - Dept. of Botany - Lab. of Plant Anatomy Porto Alegre - RS - Brazil e-mail: rinaldop-at-botanica.ufrgs.br
You might consider the advantages of a digital screen capture program. I use one called Capture for the Macintosh. It saves at the screen image resolution and allows a variety of formats (PICT, TIFF, etc.). Open it in Photoshop, clip, adjust colors, add annotation, and print on you dye-sub or ink-jet printer. No film to mess with, no chemicals, no fiddling with exposures, no wait. What could be sweeter?
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Bart, I have often photographed images on the computer screen with good results. I have found that Kodak film tends to give slides that are too blue. However Polaroid Presentation Chrome really gives you what you see. You get very good greyscale and colors are realistic as well. We use a 60mm lens and photograph from a few feet from the monitor in order to get the entire monitor screen to fill the field. However, the further away you can get from the monitor, the less your image will be affected by screen curviture. Do try to use as flat a monitor as possible. I normally will shoot an exposure series by changing the f-stop using the camera automatic exposure meter as my guide. You must have a very high resolution monitor to really produce good slides and usually an electronic slide maker will do a bit better. However, this method works well and is quite fast to do once you get the hang of setting it up. Do take the time to level the monitor so the screen is parallel with the camera, use a sturdy tripod to hold the camera and use a cable release or automatic shutter delay (if your camera is so equipped). Good luck,
Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057 --------------------------------------
I would like to photograph my multi-sync computer monitor screen using 35mm color slide film.
Assuming it is worth a try... does anyone have suggestions regarding: 1)film type 2)shutter speed 3)filters?
Thank you.
Bart Cannon Cannon Microprobe
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At 04:29 3/09/98 -0700, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
An ordinary 50 mm lens can focus close enough to use. At 1/4 second you need a tripod or some other way to steady the cameera
Its important to keep the room dark to minimise reflections from the front of the screen. Polaroid used to sell a handy plastic hood which pressed up against the screen and simultaneously eliminated reflections and steadied the camera.
We never used filters. Color is very subjective, especially when being shown as slides.
***************************************************** Mel Dickson, Director. Electron Microscope Unit, University of New South Wales. Sydney NSW 2052 Australia
On Mon, 10 Aug 1998, Dr. Gary Faulkner, Electron Microscopy Unit wrote:
} Date: Mon, 10 Aug 1998 11:47:02 AST } From: Dr. Gary Faulkner, Electron Microscopy Unit {gfaulkner-at-tupdean1.med.dal.ca} } To: Microscopy-at-sparc5.microscopy.com } Subject: EM Decline } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } While my EM Unit caters strictly to research, I am often asked } whether I would agree that the usefulness of the EM in the } clinical setting is in a major decline. It is pointed out } that diagnostic EM, especially in the area of tumors, is rapidly } being replaced by immunocytochemistry at the LM level. In addition, } tumor cells do not as a class exhibit ultrastructural changes that } can tell us much in the way of significant information anyway. I was } wondering if you might have an opinion on this observation or a } reference(s) where I could check this out. } } Many thanks, } } Gary } } gary.faulkner-at-dal.ca } I can only answer for our situation:
My Surgical Pathology EM unit is holding steady at around 350-400 cases/year--muscle, nerve, heart, kidney, tumors--all thin sections.
My EM Diagnostic Virology unit does about 1000 samples per year, steady over the last couple of years, and July-August, 1998 same as last year. This case load is mostly (90%) negative stains of fluid samples processed by negative staining; 10% is thin sectioning of tissues.
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
} I don't think 1/30th will be long enough, especially for TV's. They scan at } 60 frames per second but only refresh half the lines per pass, so it takes } 1/30th of a second to get one whole image. Now if the camera shutter is } exactly 1/30th second, then you would get one and only one complete pass. } } We tried a similar exercise on one of our EDS monitors years ago and used } too fast a speed. I think I figured we captured 4 and some scans. We could } see a brighter band on the slide where 5 frames had been scanned compared to } where only 4 frames had been scanned. } } So longer is better, and you will probably want to use a tripod for making } sure you have the same setup between shots anyway. } } Warren } } Warren,
Thanks for the correction. I was unaware of the "every other line" refresh rate, and it's a valuable piece of information. I have never had any problems photographing screens at 1/30 sec., but I haven't done this recently. Next time I'll use 1/15 or slower.
Randy
Randy Tindall 2017 Princess Jeanne Las Cruces, New Mexico 88001-4157
While on the topic of photographing monitor screens - If you focus the camera properly and focus the negative properly you can often see the scan lines quite strongly on the final print. Although it may go against the grain, slightly defocus the image on the negative and again on the print and the final image looks better for the blurring of the scan lines.
We take 0.5 and 1 sec exposures f 5.6 or 8 to get the best image from our B+W monitors.
Good luck, Ron =========================================================================== Mr. Ron Doole e-mail ron.doole-at-materials.ox.ac.uk Department of Materials, phone +44 (0) 1865 273701 University of Oxford, fax +44 (0) 1865 283333 Parks Road. Oxford. OX1 3PH. UK. ============================================================================
Rejane- Replicating with various plastic films is easy. Apply a drop or two of the recommended solvent (acetone or=20 methyl acetate) onto the surface. Cover with a small rectangle of replicating film, this=20 should have a notch near a corner to identify "specimen=20 side". Apply for a moment gentle pressure, perhaps using a=20 microscope slide. Leave to dry for five minutes. Pull the replica off. Angle coating (say 30 degrees) with metal helps to=20 visualise depths for light microscopy, whereas for SEM,=20 sputter coating is preferable. For TEM a secondary replica=20 of carbon and metal would be required, but that is another=20 story.
The replica would probably remove the plant hairs and a=20 subsequently applied replica may be preferred. Also, many=20 leaf surfaces are covered with wax. This may be removed by=20 flushing the leaf prior to replication with xylene. Rejane, if you require more details email me direct. Jim Darley ProSciTech Microscopy=20 PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au=20 *****
On Thursday, 3 September 1998 20:01, Rejane Magalh=E3es=20 Pimentel Galindo [SMTP:ggalindo-at-elogica.com.br] wrote: } } I work with Atriplex nummularia and I want to examine } epidermal cell } patterns. This specie has a proffusion of glandular } hairs in both surfaces. } } I read in a paper something about use cellulose acetate } } film with acetone } } to make impressions of the leaf. } } } } Someone know something different and simplest about=20 this } } subject? } } I also need to know what to do to take off these } } trichomes. } } Aside this, I want to obtain the vein impressions; I need } a simple method } to highlight the leaf venation. } } Thank in advance.
} Rejane Magalh=E3es Pimentel Galindo } ggalindo-at-elogica.com.br } Universidade Federal Rural de Pernambuco } Av. Boa Viagem, 6592/602 } FAX: 55 (081) 4416177 } 51130-000, Recife, Pernambuco, Brasil
Several months ago someone posted a message about difficulties they were having with quantifying synapses in LM preparations. I'd be grateful if this person would contact me offline. I've tried to find this message in the archives, without success, but that could be from my inexperience with searching there. Thanks,
I am in the market for a digital camera that can fulfill a wide range of objectives, including fluorescence imaging of fura. The experiments would probably be time-varying, which would limit exposures to ~1/2 second at best. Judging by the fact that the systems I have seen use intensified CCD (ICCD) cameras, my impression is that fura produces a very weak signal. Does anyone know if there are any non-intensified cameras (eg back-illuminated) capable of the same performance, or are ICCD cameras the only way to go?
Thanks
Eric
Eric Johnston Department of Bioengineering University of Pennsylvania 120 Hayden Hall 3320 Smith Walk Philadelphia, PA 19104-6392 215-898-1958 (F) 215-573-2071 ericdj-at-seas.upenn.edu
Prepares biological specimens for microscopic observation and analysis, using a variety of laboratory techniques and procedures. Assists in maintaining the laboratory and work environment at Bessey Microscopy Facility. Requirements: Bachelor's degree in a biological science discipline with knowledge in general chemistry, quantitative analysis, general physics, the use and general maintenance of light and electron microscopes, ancillary equipment, and all phases of handling and preparing biological specimens. Must be able to work as a part of a team, to communicate effectively with the clients, and to develop a work schedule that fits the needs of the BMF. Preferred: Master's degree in a biological science discipline; two years of laboratory experience, which may include academic courses; and practical experiences in the use of optical light microscopes, scanning and transmission electron microscopes, rotary and ultra-microtomes; darkroom experience; and knowledge of computers, networking and digital imaging; BEMT Certification (MSA). Send three (3) letters of reference and resume to Dr. Harry T. Horner, Bessey Microscopy Facility, Room 3A Bessey Hall, Iowa State University, Ames, IA 50011-1020. Fax 515.294.1337 or hth-at-iastate.edu ISU is an EO/AA employer
Tracey M. Pepper Supervisor Bessey Microscopy Facility Iowa State University Ames, IA 50011-1020 Phone: 515.294.3872 FAX: 515.294.1337 email: tpepper-at-iastate.edu
I am helping a local high school set up a scanning electron microscope for their advanced placement physics program.
Rather than draft a new course, I would like to know if anyone has a suitable teaching materials for an introduction to electron microscopy. This might include a course outline and names of textbooks.
Also, the school could use sputter coater.
Thanks in advance for help.
Alan Stone ASTON Metallurgical Services 4201 N Ravenswood Ave Chicago, IL 60613 Phone 773/528-9830
A uniform Polystyrene Latex Standard is available from SPI supplies USA, part # 02709-AB, having following specs:
Particle diameter: 0.105 um Uncertainty : 2 nm
This std. can be used to calibrate SEM. I would recommend to first calibrate the micron bar on the SEM using this std. (0.105 um particle/s). Size of the micron bar can be precisely adjusted by varying a potentiometer, provided at the display unit (with most SEM's). End result should be observed on the micrograph for comparison with actual sample. Because usually the SEM viewing screens display oversized image and have a multiplication factor to the actual magnification, while on the micrograph you get direct magnification.
Hope my 2 cents will help. Best of luck.
Zia ur Rahman E/M Engineer University of Central Florida
( Getting old is when a narrow waist and broad mind change places )
At 08:26 AM 9/4/1998 -0500, Martin Klein wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
In a message dated 98-09-04 19:17:59 EDT, you write:
{ { Martin:
A uniform Polystyrene Latex Standard is available from SPI supplies USA, part # 02709-AB, having following specs:
Particle diameter: 0.105 um Uncertainty : 2 nm } } Although the uncertainty relating to particle diameter might be 2nm one must also consider possible additional error due to shrinkage of the spheres, both during sample preparation and when subjected to high vacuum and the SEM beam.
} I am helping a local high school set up a scanning electron microscope for
} their advanced placement physics program.
}
} Rather than draft a new course, I would like to know if anyone has a
} suitable teaching materials for an introduction to electron microscopy.
} This might include a course outline and names of textbooks.
}
} Also, the school could use sputter coater.
}
} Thanks in advance for help.
}
} Alan Stone
} ASTON Metallurgical Services
} 4201 N Ravenswood Ave
} Chicago, IL 60613
} Phone 773/528-9830
Alan,
One of the best places to start re:both textbook and course materials, is "Scanning Electron Microscopy: A Student's Handbook" (Postek, et. al), available through Ladd Research Assoc, Williston VT (802-878-6711). I believe they also have a website and are probably listed at the MicroWorld metasite at www.mrwn.com.
Rejane Magalhães Pimentel Galindo wrote: =============================================== I work with Atriplex nummularia and I want to examine epidermal cell patterns. This specie has a proffusion of glandular hairs in both surfaces. I read in a paper something about use cellulose acetate film with acetone to make impressions of the leaf.
Someone know something different and simplest about this subject? I also need to know what to do to take off these trichomes.
Aside this, I want to obtain the vein impressions; I need a simple method to highlight the leaf venation. =================================================== The cellulose acetate material is generally provided by those offering EM consumables as "replicating tape" or "replicating sheets". Not all cellulose acetate is the same of course, but the grades offered from most sources tends to be acceptable for surface replication. You can find information about these materials and their use on our website.
However, the use of acetone might not be desirable since it is a solvent for the oils present on the leaf surface. Also, the dried cellulose acetate film tends to be stiff and brittle and tends to "pull off" some of the fine features from the sample making the "impression" difficult to interpret. It is also an "inverted" or "negative" image of the surface, something sometimes not so easy to interpret.
Another alternative would be our own SPI Wet Replica Kit. It was originally developed for replicating human skin in vivo and other non-dry surfaces. It too can be found on our website. It is a silicone resin based system and hence will not dissolve. It is cured with a very fast acting catalyst. The silicone really does not want to "wet" the surface and therefor the adhesion , and therefore also the tendency to pull off fine features is much less. But at this point, again you do end up with a "negative" replica.
The kit comes with another material (polyolefin) that is used to "replicate the replica" thereby generating a positive replica which should look just like the original. This now (vacuum) inert sample can be treated like any other SEM sample. This approach also makes possible the following of the same identical area, as a function of time, almost like a time-lapse- photography effect but at the SEM level.
However, like with all else in life, there are trade offs, the main one here being that above about 700X in an SEM you start to see structure from the replicating system itself. So if the features you are wanting to see can be seen below about 700X, this approach might work for you.
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
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I have been interested in FTIR microscopic spectroscopy. I have worked in IR-transmission, reflectance and absoption(?-excuse me, this mode always confuses me). Most of my work was with metals and oxides,ocasionally organics. Microscopic spectroscopy,of course, has edge and aperature shift effects. But most microscopic systems I have tried to use have seemed to have have chronic incurable alignments and operation problems. I occasionally hear a success story and several papers published, but I've most frequently heard the same chronic failure stories. Can anyone recall similar situations and a cure? Also does anyone know a list server dedicated to FTIR microscopic spectroscopy? Please address any negative info to my Email or respond generically and technically on this server. The possibilities seem very exciting in FTIR microscopic spectroscopy, but I'm afraid there is a larger group of similar problems out there. If this is the case, I'd like to work toward a cure and notify all who need and the server members. Jeff Day Mesquite, Texas Email: WA5EKH-at-Juno.Com
_____________________________________________________________________ You don't need to buy Internet access to use free Internet e-mail. Get completely free e-mail from Juno at http://www.juno.com Or call Juno at (800) 654-JUNO [654-5866]
I have been interested in FTIR microscopic spectroscopy. I have worked in IR-transmission, reflectance and absoption(?-excuse me, this mode always confuses me). Most of my work was with metals and oxides,occasionally organics. Microscopic spectroscopy,of course, has edge and aperture shift effects. But most microscopic systems I have tried to use have seemed to have had chronic incurable alignments and operation problems. I occasionally hear a success story and several papers published, but I've most frequently heard the same chronic failure stories. Can anyone recall similar situations and a cure? Also does anyone know a list server dedicated to FTIR microscopic spectroscopy? Please address any negative info to my Email or respond generically and technically on this server. The possibilities seem very exciting in FTIR microscopic spectroscopy, but I'm afraid there is a larger group of similar problems out there. If this is the case, I'd like to work toward a cure and notify all who need and the server members. Jeff Day Mesquite, Texas Email: WA5EKH-at-Juno.Com
_____________________________________________________________________ You don't need to buy Internet access to use free Internet e-mail. Get completely free e-mail from Juno at http://www.juno.com Or call Juno at (800) 654-JUNO [654-5866]
In a message dated 9/5/98 6:21:31 PM EST, cgarber-at-2spi.com writes:
} =============================================== } I work with Atriplex nummularia and I want to examine epidermal cell } patterns. This specie has a proffusion of glandular hairs in both } surfaces. I read in a paper something about use cellulose acetate film } with acetone to make impressions of the leaf.
} } Greetings. } } Does anyone know how long you can keep Formvar in solution? I need to make } } perfect films. I have a lot of Formvar in Ethylene Dichloride, but it has } } been on the shelf unopened since 1990. Can I use it? } } } } Sally Shrom } } } } } } } Hi! } Formvar solutions deterioate due to atmospheric moisture and age. This } causes holes. If not holes, then "measles" (light areas in the film). It } is not worth ruining sections because one has used old Formvar solution. } Buy or make new from powder. Make a few films and check in the TEM for } stability, thickness, etc., before making a thousand. } Bye, } Hildy } I agree with Hildy. Test it first. "the proof is in the pudding". We store formvar for up to three years so far. Normally we make small amounts an use it till it is finished. We add Molecular sieve to keep it dry and store at ~4 Deg. C to reduce evaporation.
Mr. S H Coetzee Tell: (011) 716 2419 Electron Microscope Unit Fax: (011) 339 3407 Private bag X3 E-mail: Stephan-at-gecko.biol.wits.ac.za Wits Johannesburg 2050
In his request Rejane requested help with a particular=20 replication method and I provided this. I know this method=20 works well with at least some leaves - I've done it. The hairs on these particular leaves could be a problem.=20 Possibly the best solution is to pull the hairs off with a=20 first replica and then use for microscopy a second replica,=20 which is made on the same part of the leaf. Perhaps Rejane could obtain his stated goals: identifying=20 cell patterns and vein structures, by simply using a=20 dissecting scope. Anyway, the thin plastic replica would=20 give the option to use a low power compound light=20 microscope or to use SEM. Another correspondent (also a vendor) recommended the=20 alternative of his "special" silicone double replicating=20 kit. This material may have some uses, in this case it=20 would for instance make the process more complicated and=20 more expensive. He suggested: However, the use of acetone might not be desirable since it=20 is a solvent for the oils present on the leaf surface. Also, the dried=20 cellulose acetate film tends to be stiff and brittle and tends to "pull off"=20 some of the fine features from the sample making the "impression" difficult=20 to interpret. It is also an "inverted" or "negative" image of the surface,=20 something sometimes not so easy to interpret. The questioner expressed no interest in any oils, wax does=20 not readily dissolve in acetone (though it is probably=20 desirable to dissolve any waxes to see the features of=20 interest). I have not found such replicas stiff and it would be=20 desirable to remove the hairs to expose features. If the hairs were of interest, surely routine SEM of the=20 leaf should be the recommendation. The silicone replica is of no use under a compound=20 microscope. True the replica would be a negative, but why make a=20 positive for SEM (for TEM its another story)? SEMs have a polarity switch and a flick of that switch=20 turns a negative image of a single stage replica into a=20 positive image. A non-digital photograph, taken with a light microscope=20 could be reversed by making a contact print onto a bit of=20 TEM 4489 film to achieve a negative (therefore a positive=20 of the replica) on the paper print.
Some questioners are inexperienced and are not in a=20 position to properly evaluate given advice. I prefer to recommend a product only if convinced that this=20 would aid the work of the inquirer. As a vendor I may be=20 keen to make certain products known, but that should be at=20 my expense, eg advertising. It is not fair to use the=20 Listserver feature 'MY' products, unless its quite relevant=20 to the inquiry. Its abominable, to recommend an unsuitable=20 product when in the process an inexperienced person maybe=20 misled into using an unsuitable technique and a more=20 expensive product. True, we all have an ability to=20 'convince' ourselves that our actions are 'right', but I=20 think that most people can see the difference between=20 rationalising and self-delusion. Cheers Jim Darley ProSciTech Microscopy=20 PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au=20 *****
Rejane- Replicating with various plastic films is easy. Apply a drop or two of the recommended solvent (acetone or methyl acetate) onto the surface. Cover with a small rectangle of replicating film, this should have a notch near a corner to identify "specimen side". Apply for a moment gentle pressure, perhaps using a microscope slide. Leave to dry for five minutes. Pull the replica off. Angle coating (say 30 degrees) with metal helps to visualise depths for light microscopy, whereas for SEM, sputter coating is preferable. For TEM a secondary replica of carbon and metal would be required, but that is another story.
The replica would probably remove the plant hairs and a subsequently applied replica may be preferred. Also, many leaf surfaces are covered with wax. This may be removed by flushing the leaf prior to replication with xylene. Rejane, if you require more details email me direct. Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au *****
On Thursday, 3 September 1998 20:01, Rejane Magalh=E3es Pimentel Galindo [SMTP:ggalindo-at-elogica.com.br] wrote: } } I work with Atriplex nummularia and I want to examine } epidermal cell } patterns. This specie has a proffusion of glandular } hairs in both surfaces. } } I read in a paper something about use cellulose acetate } } film with acetone } } to make impressions of the leaf. } } } } Someone know something different and simplest about this } } subject? } } I also need to know what to do to take off these } } trichomes. } } Aside this, I want to obtain the vein impressions; I need } a simple method } to highlight the leaf venation. } } Thank in advance.
26th SMG Symposium at the West Park Conference centre, Univerity Dundee. Wednesday 11 November 98
1998 Final Programme 'Apoptosis' - when discretion is better than valour David Harrison, Dept of Pathology, University of Edinburgh.
'En Bloc' optical staining of resin embedde specimens using a confocal laser scanning microscope Ian Roberts, Scottish Crop Research Institute, Dundee
Cathodoluminescence microscopy Adrian Finch, Dept Environmental Sciences, University of Hertfordshire.
Blackcurrant fruit development in three dimensions by NMR microscopy and complimentary techniques Shelia Glidewell, Scottish Crop Research Institute, Dundee
Cryo Techniques and high resolution conventional SEM Alan Robbins, Oxford Instruments, Oxford
Quantitative immunoelectron microscopy - insights into mitotic membrane dynamics John Lucocq, Department Anatomy and Physiology, University of Dundee.
Microscopy in studies of plant surface, structure and function Chris Jeffree, Dept of Botany, University of Edinburgh.
Visit our Web Site at: http://www.abdn.ac.uk/~nhi691/smg98.htm
EM Unit, Dept Zoology University Of Aberdeen Tillydrone Avenue Aberdeen AB24 2TZ Tel 01224-272847 Fax 01224-272396
recently there was a discussion about scanners. If I'm remember well the Leaf scanner is now produce by a compagny named Brenson Inc. . Do some of you now these compagny and have they a sale representative in Europe ?
Thank's a lot
Marc
------------------------------ SCHMUTZ Marc IGBMC 1 rue Laurent FRIES BP 163 F 67404 Illkirch Cedex FRANCE
Here's a good trick. I recently wanted to do leaf surface impressions for stomatal counts and was experimenting with cellulose acetate etc. A physiologist friend comes along and says "have you tried ignition sealer?" Turns out this works really well. I sprayed the leaf surface with Kraco ignition sealant (this stuff is used as a spray waterproofing treatment for electrical wires and engine components), let it set for 5 min. and then peeled off the surface coating with clear tape. The replica/tape was then simply stuck to a glass slide and examined by phase contrast. This works well for epidermal cell shape, stomatal shape and distribution, and vein shape. I don't know how well it will work on trichomes and it probably depends on how elaborated they are. A word of caution; it seems that not all ignition sealer sprays are created equal. 'Wire Dryer' brand did not work at all. Other companies that manufacture this stuff include Hydrosol, Kleenflo and Spray-pak.
} } } =============================================== } } I work with Atriplex nummularia and I want to examine epidermal cell } } patterns. This specie has a proffusion of glandular hairs in both } } surfaces. I read in a paper something about use cellulose acetate film } } with acetone to make impressions of the leaf. } } How about gelatin? } } Jim Harper
________________________ C. John Runions, Ph.D. Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
Hello all, I've recently had a request to do some TEM of some PbSn solder bumps (on silicon ICs). I have some concerns about this, since the solder can begin to melt at about 150C. Has anyone been sucessful in making a TEM section of such a structure? I would be very interested in any tips or tricks.
Also, our local supplier of glassine envelopes for TEM negatives (3 3/4 x 2 3/4 inch) has told us they aren't stocking any more. Does anyone know of a UK-based supplier?
Many thanks in advance
Richard Beanland GMMT Ltd., Caswell, Towcester, Northants NN12 8EQ UK
Hi all, in my earlier email about spraying ignition sealer onto leaves to make replicas I said that the product used was manufactured by Kraco. I lied. It's actually made by Krylon of Columbus Ohio, 43215. Cheers, John
________________________ C. John Runions, Ph.D. Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
} I've recently had a request to do some TEM of some PbSn solder bumps } (on silicon ICs). I have some concerns about this, since the solder can begin } to melt at about 150C. Has anyone been sucessful in making a TEM section of } such a structure? I would be very interested in any tips or tricks. } I suggest examining the specimen at ~-150 C. You do not need to cool the specimen prior to insertion in the TEM; just place the RT grid in the cryo holder, insert the holder into the TEM, then cool. Low dose rate will also help keep beam heating from producing locally high temperatures. Good luck. Yours, Bill Tivol
I am in the market for a digital camera that can fulfill a wide range of objectives, including fluorescence imaging of fura. The experiments would probably be time-varying, which would limit exposures to ~1/2 second at best. Judging by the fact that the systems I have seen use intensified CCD (ICCD) cameras, my impression is that fura produces a very weak signal. Does anyone know if there are any non-intensified cameras (eg back-illuminated) capable of the same performance, or are ICCD cameras the only way to go?
Thanks
Eric
Eric Johnston Department of Bioengineering University of Pennsylvania 120 Hayden Hall 3320 Smith Walk Philadelphia, PA 19104-6392 215-898-1958 (F) 215-573-2071 ericdj-at-seas.upenn.edu
I'm new at operating an SEM that has a tungsten filament. For the past 5 years I had an instrument that used a LaB6, but I've changed companies. I have 2 questions. 1. Both the people here and the instrument's service man tell me that the tungsten filament is more stable than the LaB6. I ask them to explain further and they really don't get into it. What is the stable thing about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, so what is the unstableness of it? 2. I'm also told here that, with the tungsten filament, as the beam sits in one area on the sample, Carbon will develop in that area. Is this true? If so how long does it take for the carbon to contaminate the area, and also does this take any confidence in a carbon analysis and throw it out the window?
For a brief explanation check J.W. Edington. "Practical Electron Microscopy in Materials Science"
or you can check Hirsch et.al. "Electron Microscopy of Thin Crystals" Chapter 16.
Very briefly, Lorentz Microscopy is a technique used to image magnetic domains in ferromagnetic materials using a TEM. The name comes from the Lorentz formula ( i.e. v xB see an introductory Physics book) for the force on a charged particle moving in a magnetic field B.
Jordi Marti ---------- } From: "Pettieswa-at-aol.com"-at-sparc5.microscopy.com To: Microscopy-at-sparc5.microscopy.com -----------------------------------------------------------------------
In a message dated 98-09-08 09:24:58 EDT, ericdj-at-seas.upenn.edu writes:
{ { I am in the market for a digital camera that can fulfill a wide range of objectives, including fluorescence imaging of fura. The experiments would probably be time- varying, which would limit exposures to ~1/2 second at best. Judging by the fact that the systems I have seen use intensified CCD (ICCD) cameras, my impression is that fura produces a very weak signal. Does anyone know if there are any non-intensified cameras (eg back- illuminated) capable of the same performance, or are ICCD cameras the only way to go? } }
Eric,
I'm not sure what a "back-illuminated non-intensified" camera is, but I would highly recommend a cooled CCD camera for your applications, probably something like the Spot or maybe the Spot Jr., from Diagnostic Instruments ($5-8K).
It depends on your application, but a 1/2 second exposure would be pushing it for just about any CCD camera if you want color. There are other high-end, high-speed digital cameras (maybe something from Dage MTI) that could do this in B&W, but they are in the range of $20-25K.
Hope this is of some help.
Bob ********************************* Robert (Bob) Chiovetti E. Licht Company / 1-800-865-4248 / (520) 546-4986 rchiovetti-at-aol.com
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At 10:37 AM 9/8/98 -0400, Mark Darus wrote: } } I'm new at operating an SEM that has a tungsten filament. For the past 5 } years I had an instrument that used a LaB6, but I've changed companies. } I have 2 questions. } 1. Both the people here and the instrument's service man tell me } that the tungsten filament is more stable than the LaB6. I ask them to } explain further and they really don't get into it. What is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, } so what is the unstableness of it?
I have only ever run tungsten, but the current has been stable for me. I can't say anything about instabilities. I have heard about field emission scopes sometimes being unstable, but even that has remedies.
} 2. I'm also told here that, with the tungsten filament, as the beam sits in } one area on the sample, Carbon will develop in that area. Is this true? If } so how long does it take for the carbon to contaminate the area, and also } does this take any confidence in a carbon analysis and throw it out the } window?
Last I heard, there was no difference between electrons once they left the gun, be it W, LaB6, or FE. The only difference would be in the number of them per time in a given space. The lesser vacuum requirements for a W filament might lead to more rapid C buildup on the sample. But our 840 with W normally runs 10-6 torr and has only moderate problems with buildup.
I am leary of doing much with C analyses. The absorption factors are significant sources of uncertainty. I would stick to qualitative comparisons between points as much as possible. And make sure spectra are collected under similar conditions of time and current.
Dear Mark, } } 1. Both the people here and the instrument's service man tell me } that the tungsten filament is more stable than the LaB6. I ask them to } explain further and they really don't get into it. What is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, } so what is the unstableness of it?
Can you put a Faraday cage in the beam? If so, you can measure the stability for periods longer than the response time of the cage+electronics. High-frequency instabilities are harder to measure, but would show up as light or dark bands in the image for appropriate scan rates. I have no experience with SEM, but I don't think there is much difference in stabil- ity between LaB6 and W. Both are thermionic sources which should be heated by DC current. In this case the temperature--therefore the emission--should be constant.
} 2. I'm also told here that, with the tungsten filament, as the beam sits in } one area on the sample, Carbon will develop in that area. Is this true? If } so how long does it take for the carbon to contaminate the area, and also } does this take any confidence in a carbon analysis and throw it out the } window? } This will happen with any source of electrons, but the contamination rate will vary with such parameters as vacuum in the specimen area and the nature of the specimen. The appearance of these contaminant peaks (they look like little mountains) has been used to measure local specimen thickness and to identify the position where EDS was done (for single-point analysis). The HVEM does not leave such peaks in spite of the so-so column vacuum and its use for plastic sections of biological specimens. Certainly, if you see carbon peaks on the specimen, you will also see carbon peaks in the EDS spec- tra, so I wouldn't trust a carbon analysis on such an instrument either for a single-point spectrum or for a carbon element map. Yours, Bill Tivol
I am looking for pointers to products or articles that address the following idea.
A single picture taken through a light microscope at say 100x has very small depth of focus.
But if a large number of pictures is taken at varying focal planes, the entire surface of a 3D subject can be imaged sharply, albeit one plane at a time.
It seems, in concept, that the sharply focused portions of each picture could be montaged together, to produce a single picture showing the entire 3D surface in sharp focus at once.
The required number of pictures could be quite large, but the montaging seems like a job that could be automated with computer image processing software.
I presume that this idea must have been studied somewhere. Perhaps there are products available to do the job. But I don't know them.
So I am asking for help. If you know of literature articles, products, or people who are working on this technique, then I would greatly appreciate getting a pointer to them.
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Dear Mark, } } 1. Both the people here and the instrument's service man tell me } that the tungsten filament is more stable than the LaB6. I ask them to } explain further and they really don't get into it. What is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, } so what is the unstableness of it?
Can you put a Faraday cage in the beam? If so, you can measure the stability for periods longer than the response time of the cage+electronics. High-frequency instabilities are harder to measure, but would show up as light or dark bands in the image for appropriate scan rates. I have no experience with SEM, but I don't think there is much difference in stabil- ity between LaB6 and W. Both are thermionic sources which should be heated by DC current. In this case the temperature--therefore the emission--should be constant.
} 2. I'm also told here that, with the tungsten filament, as the beam sits in } one area on the sample, Carbon will develop in that area. Is this true? If } so how long does it take for the carbon to contaminate the area, and also } does this take any confidence in a carbon analysis and throw it out the } window? } This will happen with any source of electrons, but the contamination rate will vary with such parameters as vacuum in the specimen area and the nature of the specimen. The appearance of these contaminant peaks (they look like little mountains) has been used to measure local specimen thickness and to identify the position where EDS was done (for single-point analysis). The HVEM does not leave such peaks in spite of the so-so column vacuum and its use for plastic sections of biological specimens. Certainly, if you see carbon peaks on the specimen, you will also see carbon peaks in the EDS spec- tra, so I wouldn't trust a carbon analysis on such an instrument either for a single-point spectrum or for a carbon element map. Yours, Bill Tivol
I would check with Diagnostic Instruments, Inc. Sterling Heights, Michigan 313-731-6000. There is probably a dealer in your area, but am not sure who.
Good Luck, C. Passione -----Original Message----- } From: Eric Johnston {ericdj-at-seas.upenn.edu} To: 'Microscopy' {Microscopy-at-sparc5.microscopy.com}
Thank you for all your responses to my need for a critical point dryer. I have purchased one, but have saved the information (vendors, models, prices, etc.) from the other offers I received. If anyone else is looking for a good price on a CPD, drop me a line, and I'll share the info.
The Joint Meeting of the Florida American Vacuum Society and the Florida Society for Microscsopy will be held the week following PITTCON (March 14-16, 1999) in Orlando on the campus of the University of Central Florida.
Events
golf tournament 2 day symposium (physical and biological sciences) including over 20 invited speakers student poster session and prizes: 1st prize: all expenses paid trip to attend national AVS or MSA meetings over 40 exhibitors AVS sponsored short courses equipment demos
AND Vendor Sponsored short courses:
1) Specimen Preparation using the Tripod Polisher (3/12-3/13): sponsored by South Bay Technology, instructor: Ron Anderson, IBM.
2) Specimen Preparation using Focused Ion Beam Milling (3/18-3/19): sponsored by FEI/Philips, instructors: Lucille Giannuzzi, UCF and Fred Stevie, Cirent Semiconductor.
Vendor space is limited and sells out early. Please contact Fred Stevie at stevie-at-lucent.com.
For other information, please contact Lucille Giannuzzi at lag-at-pegasus.cc.ucf.edu
******************************************************************* Lucille A. Giannuzzi, Ph.D.
Associate Professor, Dept. of Mechanical, Materials, and Aerospace Eng., University of Central Florida, PO Box 162450, 4000 Central Florida Blvd., Orlando, FL 32816-2450 USA phone (407) 823-5770 fax (407) 823-0208 email lag-at-pegasus.cc.ucf.edu
Director, UCF/Cirent Materials Characterization Facility, 12443 Research Parkway, Suite 305 Orlando, FL 32826 phone (407) 275-4354,5,6 fax (407) 275-4321 -------------------------------------------------------------------- "Good judgement comes from experience.
Experience comes from making bad judgement."
Mark Twain ********************************************************************
} } So I am asking for help. If you know of literature articles, } products, or people who are working on this technique, then I would } greatly appreciate getting a pointer to them. } } Please respond by email to } } Rik.Littlefield-at-pnl.gov (or rj_littlefield-at-pnl.gov) } } Thanks very much. } =================================================================== } Rik Littlefield } Senior Research Scientist } Pacific Northwest National Laboratory } Richland, WA 99352 } email: Rik.Littlefield-at-pnl.gov } phone: 509-375-3927 } fax: 509-375-3641
Hi,
Lots of information available for your interest, I'm deep into this kinda work. Since I work with 3-d reconstructions of injured spinal cord tissue and transplants and other things (you?) I'll generalize.
1. About sectioning as you mentioned, this can be done on thick sections via "optical sectioning" using a microscope with z-axis motor (Leica, for example), ours is computer driven with a CCD camera to automatically acquire a "stack" of images through thick tissue sections. However, depending on what you're considering, using physically sectioned (histological section) with embedded fiducial points for section registration can also be quite useful (lots of literature on that), thereafter, fairly large histological sections (say 25mm long by 15 mm wide) can be imaged under high resolution (less than 10 microns per pixel) which can allow reconstructions where optical section might not work (for large tissue regions). Course you won't get the resolution of 100x. IPLab by Scanalytics is a good program that might be helpful (forgot their URL).
2. You can take sections that are "in focus" based on distinct boundaries that can be automatically identified via algorithm. The freeware image program NIH-Image (http://rsb.info.nih.gov/nih-image/) with one of the myriad of already available macros (forgot the name of the specific one but you can do a gopher search on the NIH-Image website of the NIH-Image discussion list) can do this with a stack of images to create a composite (aside, I think you mean "composite" versus "montage" since we're dealing with 3-d and not strictly 2-d imaging). Oh yeah, if you are into imaging, get yourself on that NIH-Image discussion list. You'll meet other imaging experts there.
3. Also, there are programs such as Surfdriver (http://surfdriver.ml.org) which you can use to outline boundaries on a stack of images and create shaded surfaces of 3-d structures. Going deeper, you can check out VoxBlast (http://www.vaytek.com/) which does more sophisticated 3-D volume visualization and allows measurements and other interesting things (handles my 300-400 megabyte datasets like a charm). You don't mention digital deconvolution so we won't go there. Going deeper still, why go to all the trouble to reconstruct something in 3-D, without thinking spatial analysis? Perhaps, you want to know how close a dendrite surface is to a particular neuron and would like a 3-D color surface map (like a terrain map) detailing proximity (or it could be 3-D localized concentrations of transmitter or whatever). For that kind of custom work, check out IDL from RSI, Inc. (http://www.rsinc.com/) but now we're into programming. Point is, lots o' people want to visualize in 3-D, but the fun begins when you want to quantify in 3-D. Oh, and if you quantify, think "stereology."
For literature searches, I would suggest "3-D reconstruction" (text search) on Medline as a start and of course, check out the latest book (3rd edition) of John Russ's "The Image Processing Handbook" by CRC Press, and is just fabulous (unpaid endorsement) plus he has a way-cool course he teaches in North Carolina which I thought was great, plus he has way-cool software CD-ROM Image Processing Toolkit (http://members.aol.com/ImagProcTK/) with useful tutorials, very good. Not heavy into 3-D but the 2-D coverage is so useful you should just have it.
Well, enough gushing over 3-D and imaging, back to work for me, hope I said something useful. Brian C. Tryon MD/PhD student Allegheny University of the Health Sciences (Hey! We're bankrupt!) School of Medicine 3200 Henry Avenue Philadelphia, PA 19129 USA
I can speak from experience here since I have been using LaB6 & Tungsten for a number of years.
Tungsten in general will give a short lifetime ( typically 100 hrs+ ) whilst a LaB6 ( in theory ) will last for months. My experience with LaB6, which probably depends on the LaB6 supplier ( in this case Denka ), has produced approx. 6 weeks of more electrons before the filament starts to produce less and the alignment wander.
This may be due to the type of source of the LaB6, but I have talked recently to someone running a different supplier's LaB6 and he was suffering the same problems.
In general operation the LaB6 will not give noticable differences in stability. Any measurements using a Faraday cage & monitor will give different results depending on the age of the filament. However if you have demanding customers, such as Geologists ( apologies to any reading this ! ), when you are carrying out quant analyses and producing totals approaching 100% the analyses are very dependant on beam stability. I have not been able to carry out these analyses without using a tungsten source. Even with a tungsten filament it will be unstable in the early & late part of its life and serious analyses can only be carried out during the middle period. I produce longer life and greater stability by cleaning the gun properly after a blown filament and using the Ion pump to produce a higher vacuum during tungsten operation. Tungsten filaments also have faster ramp times and are more tolerant of poor vacuums.
As regards carbon build up I think that it is both vacuum & sample dependant. With more hydrocarbons in the vacuum you get a greater build-up. I have two turbo-molecular pumped systems and do not suffer from much carbon build-up unless certain samples are scanned.
Best wishes,
Colin
Colin Reid, Electron Microscope Unit, Trinity College Dublin, Dublin 2, Ireland. Tel: 353-1-6081820 Fax: 353-1-6770438 email: creid-at-tcd.ie -----Original Message----- } From: Mark Darus {DARUSM-at-cle.lg.bfg.com} To: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com}
Although I am a "vendor", allow me to comment on this thread, from the perspective of someone who has manufactured tungsten filaments and distributed Denka LaB6 cathodes for more than 10 years.
Colin Reid wrote:
} Tungsten in general will give a short lifetime ( typically 100 hrs+ ) whilst } a LaB6 ( in theory ) will last for months. My experience with LaB6, which } probably depends on the LaB6 supplier ( in this case Denka ), has produced } approx. 6 weeks of more electrons before the filament starts to produce less } and the alignment wander. This may be due to the type of source of the LaB6, but I have } talked recently to someone running a different supplier's LaB6 and he was suffering } the same problems.
For any filament, by far the most important parameter affecting material loss, and, therefore, lifetime is vacuum. This holds true for both tungsten filaments and LaB6. Filament life, therefore, is highly dependent on the vacuum conditions of a given microscope, and filament life will vary substantially from microscope to microscope.
Tungsten filaments are cold-formed from wire, and the stress from the forming will cause instability if the filaments are not properly annealed under vacuum. If the tungsten filament has not been properly annealed by the manufacturer, the user will, in effect, anneal it in the microscope. The filament will be unstable during this process, and will often have to be realigned after the annealing.
In the case of LaB6, the crystal is in {100} orientation, and formed into a point, a round tip or a microflat tip. As the cathode experiences material loss, the tip flattens. After some 150 hours, the brightness will fall off, as the emission surface of the tip forms into a larger flat. In effect, the sharp or round tips become flat tips. Although the cathode has hundreds of hours of "life" left, it will never again be as bright as it was during the first ~150 hours. This is true of any manufacturer's LaB6.
For quantitative analysis, mechanical stability becomes an issue. For this application, the LaB6 mount should be as robust as possible.
I would be happy to provide additional details to anyone interested. In particular, we have several Denka technical reports which explain the relationship between lifetime, brightness, vacuum and material loss for LaB6 cathodes.
Best regards, Steven E. Slap, Vice-President
******************************** Energy Beam Sciences, Inc. The Laboratory Microwave Company http://www.ebsciences.com ********************************
I saw a demo in London of a software package "Auto-Montage" from Synoptics which was combining 5 and more partially focussed images to one fully focused one, using a frame-grabber. E-mail address : sales-at-synoptics.co.uk With best regards, emond de Roever
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I am looking for pointers to products or articles that address the following idea.
A single picture taken through a light microscope at say 100x has very small depth of focus.
But if a large number of pictures is taken at varying focal planes, the entire surface of a 3D subject can be imaged sharply, albeit one plane at a time.
It seems, in concept, that the sharply focused portions of each picture could be montaged together, to produce a single picture showing the entire 3D surface in sharp focus at once.
The required number of pictures could be quite large, but the montaging seems like a job that could be automated with computer image processing software.
I presume that this idea must have been studied somewhere. Perhaps there are products available to do the job. But I don't know them.
So I am asking for help. If you know of literature articles, products, or people who are working on this technique, then I would greatly appreciate getting a pointer to them.
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Hi Mark, } } 1. Both the people here and the instrument's service man tell me } that the tungsten filament is more stable than the LaB6. I ask them to } explain further and they really don't get into it. What is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, } so what is the unstableness of it?
I have experiences on both tungsten and field emission SEMs. The emission of cold FE gun does have fluctuation. But at normal condition, the fluctuation is tolerable and can't be seen on the final images.
} 2. I'm also told here that, with the tungsten filament, as the beam sits in } one area on the sample, Carbon will develop in that area. Is this true? If } so how long does it take for the carbon to contaminate the area, and also } does this take any confidence in a carbon analysis and throw it out the } window? }
The carbon build-up, mostly hydrocarbon, is depended on vacuum and sample. Especially the "cleanliness" of the vacuum. I have worked on two different FESEMs, both are equipped with turbo pumps. The difference is that one scope is backed by a rotary pump, the other is a totally oil-free pumping system. I looked the same test sample using both scopes and found that there is a big difference on the contamination rate. I can multiple-scan the same area at the magnification of 500,000x using the FESEM equipped with totally oil-free pumping system, but not the other one.
Ya Chen
======================================================================== \ / Integrated Microscopy Resource (IMR)-- \ / __ a NIH Biomedical Research Resource TEL: 608-263-8481 \/ / / University of Wisconsin-Madison FAX: 608-265-4076 / / / 1675 Observatory Drive #159 / /__/_ Madison, WI 53706 Email: ychen14-at-facstaff.wisc.edu ======================================================================== IMR WWW Home Page: http://www.bocklabs.wisc.edu/imr/home.htm
You have accurately described a technique which is solved using several different approaches.
The most common commercialized version is confocal microscopy. The best general reference is the {underline} Handbook of Biological Confocal Microscopy {/underline} , published by Plenum and edited by Dr. James Pawley. We also have a short discussion in our book, {underline} Optimizing Light Microscopy for Biological and Clinical Laboratories {/underline} (see our web site: { {http://www.MME-Microscopy.com/education} ). You may also want to visit the web sites of the confocal manufacturers. The best metasites for locating those URLs are either the MSA manufacturers' list or MicroWorld resources and news (www.mwrn.com).
A scond commercialized approach is the 3-D imaging offered through Edge Scientific. I believe their website is listed on both the MSA and mwrn sites. They provide a variety of options, ranging from a full blown system (the R400), to a retrofittable reflected light/fluorescence head and the new "Perceptra", a retrofittable, 3-D illuminator.
I have also heard of non-commercial work done by people such as Dr. Shinya Inoue (retired from the Woods Hole Biological Institute) who have used DIC to take very thin optical sections then used computer programs to reconstruct the resulting set of serial optical sections.
There is a company that does exactly what you are describing. The company is called Soft Imaging system Corporation. I believe, they are in Denver. I know they have a website, but I can't remember the URL right now. I'll try to find out and send it to you. Perhaps somebody else knows it.
Gloria
} } } ---------- } From: Rik Littlefield[SMTP:rj_littlefield-at-pnl.gov] } Sent: Tuesday, September 8, 1998 1:02 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Photomontaging for increased depth of field ? } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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There is a company that does exactly what you are describing. The company is called Soft Imaging system Corporation. I believe, they are in Denver. I know they have a website, but I can't remember the URL right now. I'll try to find out and send it to you. Perhaps somebody else knows it.
Gloria
} } } ---------- } From: Rik Littlefield[SMTP:rj_littlefield-at-pnl.gov] } Sent: Tuesday, September 8, 1998 1:02 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Photomontaging for increased depth of field ? } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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Hi Mark, } } 1. Both the people here and the instrument's service man tell me } that the tungsten filament is more stable than the LaB6. I ask them to } explain further and they really don't get into it. What is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, } so what is the unstableness of it?
I have experiences on both tungsten and field emission SEMs. The emission of cold FE gun does have fluctuation. But at normal condition, the fluctuation is tolerable and can't be seen on the final images.
} 2. I'm also told here that, with the tungsten filament, as the beam sits in } one area on the sample, Carbon will develop in that area. Is this true? If } so how long does it take for the carbon to contaminate the area, and also } does this take any confidence in a carbon analysis and throw it out the } window? }
The carbon build-up, mostly hydrocarbon, is depended on vacuum and sample. Especially the "cleanliness" of the vacuum. I have worked on two different FESEMs, both are equipped with turbo pumps. The difference is that one scope is backed by a rotary pump, the other is a totally oil-free pumping system. I looked the same test sample using both scopes and found that there is a big difference on the contamination rate. I can multiple-scan the same area at the magnification of 500,000x using the FESEM equipped with totally oil-free pumping system, but not the other one.
Ya Chen
======================================================================== \ / Integrated Microscopy Resource (IMR)-- \ / __ a NIH Biomedical Research Resource TEL: 608-263-8481 \/ / / University of Wisconsin-Madison FAX: 608-265-4076 / / / 1675 Observatory Drive #159 / /__/_ Madison, WI 53706 Email: ychen14-at-facstaff.wisc.edu ======================================================================== IMR WWW Home Page: http://www.bocklabs.wisc.edu/imr/home.htm
Gut tissue fixation I am looking for a protocol for intestinal tissue. I have opossum intestinal tissue fixed by a colleague that has a lot of OsO4 pepper. I have also received gut tissue that appeared to have undergone autolysis. I am expecting more of these samples and would like to recommend a change in protocol. It is probably not possible to use perfusion. I am looking for intercellular and intracellular bacteria and /or protozoal infections in these tissues.=20 Would the chemistry of the intestine be causing this type of problem? Or is it simply a matter of inadequate washing?=20 Thanks=85=85.. Sally
The fixation used was:=20 2% glut in PBS at pH 7.2 for 3 hours. 3X washed in PBS 1X washed in H20 1% OsO4 for 4H room T washed 3x in H20 dehydrated in a 25% series of acetone Infiltrated and embedded in a 25% series into a Spurrs/ Quetol blend.
Sally Burns Center for Electron Optics B5 Pesticide Research Center (517) 355-5004
We have a Hitachi S-570 SEM and we would like to take "motion pictures" of MEMS devices inside the chamber using the SEM. It does have a "video out" connector on its pc board, however, the signal from it is not a standard NTSC video signal. I've talked to Hitachi and was told that at one time they made a coverter, but that it is no longer available. Does anyone, by any chance, know if the video signal conforms to some other video standard? Also, does anyone know of a source for either the Hitachi converters or some other converter that would give us a usable video signal to feed into monitor or video recorder?
Janet Rice MCC Senior Member Technical Staff rice-at-mcc.com 512-338-3266
Hi Mark, } } 1. Both the people here and the instrument's service man tell me } that the tungsten filament is more stable than the LaB6. I ask them to } explain further and they really don't get into it. What is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, perfect in fact, } so what is the unstableness of it?
I have experiences on both tungsten and field emission SEMs. The emission of cold FE gun does have fluctuation. But at normal condition, the fluctuation is tolerable and can't be seen on the final images.
} 2. I'm also told here that, with the tungsten filament, as the beam sits in } one area on the sample, Carbon will develop in that area. Is this true? If } so how long does it take for the carbon to contaminate the area, and also } does this take any confidence in a carbon analysis and throw it out the } window? }
The carbon build-up, mostly hydrocarbon, is depended on vacuum and sample. Especially the "cleanliness" of the vacuum. I have worked on two different FESEMs, both are equipped with turbo pumps. The difference is that one scope is backed by a rotary pump, the other is a totally oil-free pumping system. I looked the same test sample using both scopes and found that there is a big difference on the contamination rate. I can multiple-scan the same area at the magnification of 500,000x using the FESEM equipped with totally oil-free pumping system, but not the other one.
Ya Chen
======================================================================== \ / Integrated Microscopy Resource (IMR)-- \ / __ a NIH Biomedical Research Resource TEL: 608-263-8481 \/ / / University of Wisconsin-Madison FAX: 608-265-4076 / / / 1675 Observatory Drive #159 / /__/_ Madison, WI 53706 Email: ychen14-at-facstaff.wisc.edu ======================================================================== IMR WWW Home Page: http://www.bocklabs.wisc.edu/imr/home.htm
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Date 9/9/98 Time: 3:54 PM Internal Memorandum
We currently have two positions open for SEM/EDS materials technicians at Exponent Failure Analysis Associates in Menlo Park, CA. See attachment for details and send resume/information to hr-at-exponent.com.
by arwen.otago.ac.nz (8.9.1/8.9.1) with ESMTP id NAA17599; Thu, 10 Sep 1998 13:21:44 +1200 (NZST) X-Sender: st004716-at-brandywine.otago.ac.nz Message-Id: {l03130302b21cdafa0cf4-at-[139.80.34.56]} In-Reply-To: {3.0.32.19980909161244.0090e6d0-at-pilot.msu.edu} Mime-Version: 1.0 Content-Type: text/plain; charset="iso-8859-1" Content-Transfer-Encoding: quoted-printable
} Gut tissue fixation } I am looking for a protocol for intestinal tissue. I have opossum } intestinal tissue fixed by a colleague that has a lot of OsO4 pepper. I } have also received gut tissue that appeared to have undergone autolysis. I } am expecting more of these samples and would like to recommend a change in } protocol. It is probably not possible to use perfusion. } I am looking for intercellular and intracellular bacteria and /or } protozoal infections in these tissues. } Would the chemistry of the intestine be causing this type of } problem? Or } is it simply a matter of inadequate washing? } Thanks=85=85.. Sally } } The fixation used was: } 2% glut in PBS at pH 7.2 for 3 hours. } 3X washed in PBS } 1X washed in H20 } 1% OsO4 for 4H room T } washed 3x in H20 } dehydrated in a 25% series of acetone } Infiltrated and embedded in a 25% series into a Spurrs/ Quetol blend.
Sally, Are you sure that is OsO4 pepper? Could it be PO4 pepper? Although it seems that there are heaps of water washes in your protocol anyway, so possibly may not be. We have processed this stuff before, using perfusion admitedly, but using Cacodylate buffer, havent had too many probs as far as I can remember. I would change buffer initially to have a go. Is it necessary to fix for 4hrs? We routinely fix in 1%OsO4 in buffer for 1hr at RT. Perhaps having it in here for too long would cause excess OsO4 deposition. About the apparent autolysis, could it be that there is lots of mucus/garbage around the tissue preventing fix from getting in? Would a paraformaldehyde/glut fix be better?
Just thinking aloud.............! Hope you find the cure to your problems,
Rich.
----------------------------------------------------------------------- Richard Lander Electron Microscope Technician South Campus Electron Microscope Unit Otago School of Medical Sciences P.O. Box 913 Dunedin New Zealand. Tel. National 03 479 7301 Fax. National 03 479 7254
"Southernmost EM Unit in the World!" ------------------------------------------------------------------------
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=3EGut tissue fixation =3E I am looking for a protocol for intestinal tissue=2E I have = opossum =3Eintestinal tissue fixed by a colleague that has a lot of OsO4 pepper= =2E I =3Ehave also received gut tissue that appeared to have undergone autoly= sis=2E I =3Eam expecting more of these samples and would like to recommend a cha= nge in =3Eprotocol=2E It is probably not possible to use perfusion=2E =3E I am looking for intercellular and intracellular bacteria an= d /or =3Eprotozoal infections in these tissues=2E =3E Would the chemistry of the intestine be causing this type of=
=3Eproblem=3F Or =3Eis it simply a matter of inadequate washing=3F =3EThanks=85=85=2E=2E Sally =3E =3EThe fixation used was=3A =3E2% glut in PBS at pH 7=2E2 for 3 hours=2E =3E3X washed in PBS =3E1X washed in H20 =3E1% OsO4 for 4H room T =3Ewashed 3x in H20 =3Edehydrated in a 25% series of acetone =3EInfiltrated and embedded in a 25% series into a Spurrs=2F Quetol ble= nd=2E
Sally=2C Are you sure that is OsO4 pepper=3F Could it be PO4 pepper=3F Althoug= h it seems that there are heaps of water washes in your protocol anyway=2C s= o possibly may not be=2E We have processed this stuff before=2C using pe= rfusion admitedly=2C but using Cacodylate buffer=2C havent had too many probs a= s far as I can remember=2E I would change buffer initially to have a go=2E Is it necessary to fix for 4hrs=3F We routinely fix in 1%OsO4 in buffe= r for 1hr at RT=2E Perhaps having it in here for too long would cause excess= OsO4 deposition=2E About the apparent autolysis=2C could it be that there is lots of mucus=2Fgarbage around the tissue preventing fix from getting in=3F Wo= uld a paraformaldehyde=2Fglut fix be better=3F
Just thinking aloud=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E! Hope you find the cure to your problems=2C
Richard Lander Electron Microscope Technician South Campus Electron Microscope Unit Otago School of Medical Sciences P=2EO=2E Box 913 Dunedin New Zealand=2E Tel=2E National 03 479 7301 Fax=2E National 03 479 7254
=22Southernmost EM Unit in the World!=22 -----------------------------------------------------------------------= -
We are selling our 5 year old Leo(Leica) S440, with availibility approx. = 2-3 months. If anyone ( Ireland & possibly UK ) is interested will you = contact me directly for full details.
Thanks,
Colin
Colin Reid, Electron Microscope Unit, Trinity College Dublin, Dublin 2, Ireland. Tel: 353-1-6081820 Fax: 353-1-6770438 email: creid-at-tcd.ie
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=3EGut tissue fixation =3E I am looking for a protocol for intestinal tissue=2E I have = opossum =3Eintestinal tissue fixed by a colleague that has a lot of OsO4 pepper= =2E I =3Ehave also received gut tissue that appeared to have undergone autoly= sis=2E I =3Eam expecting more of these samples and would like to recommend a cha= nge in =3Eprotocol=2E It is probably not possible to use perfusion=2E =3E I am looking for intercellular and intracellular bacteria an= d /or =3Eprotozoal infections in these tissues=2E =3E Would the chemistry of the intestine be causing this type of=
=3Eproblem=3F Or =3Eis it simply a matter of inadequate washing=3F =3EThanks=85=85=2E=2E Sally =3E =3EThe fixation used was=3A =3E2% glut in PBS at pH 7=2E2 for 3 hours=2E =3E3X washed in PBS =3E1X washed in H20 =3E1% OsO4 for 4H room T =3Ewashed 3x in H20 =3Edehydrated in a 25% series of acetone =3EInfiltrated and embedded in a 25% series into a Spurrs=2F Quetol ble= nd=2E
Sally=2C Are you sure that is OsO4 pepper=3F Could it be PO4 pepper=3F Althoug= h it seems that there are heaps of water washes in your protocol anyway=2C s= o possibly may not be=2E We have processed this stuff before=2C using pe= rfusion admitedly=2C but using Cacodylate buffer=2C havent had too many probs a= s far as I can remember=2E I would change buffer initially to have a go=2E Is it necessary to fix for 4hrs=3F We routinely fix in 1%OsO4 in buffe= r for 1hr at RT=2E Perhaps having it in here for too long would cause excess= OsO4 deposition=2E About the apparent autolysis=2C could it be that there is lots of mucus=2Fgarbage around the tissue preventing fix from getting in=3F Wo= uld a paraformaldehyde=2Fglut fix be better=3F
Just thinking aloud=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E=2E! Hope you find the cure to your problems=2C
Richard Lander Electron Microscope Technician South Campus Electron Microscope Unit Otago School of Medical Sciences P=2EO=2E Box 913 Dunedin New Zealand=2E Tel=2E National 03 479 7301 Fax=2E National 03 479 7254
=22Southernmost EM Unit in the World!=22 -----------------------------------------------------------------------= -
May I introduce you to the Volumetric Image Processing, Analysis and Visualization software package, EIKONA3D for Windows 95 and NT that was specifically designed for 3D microscopy. This software package has been built for processing, analyzing and visualizing microscopy volumes (3D data sets or series of slices).
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If you want to receive the free demo by email, please reply to this message or contact us at alphatec-at-alphatecltd.com
Furthermore, we can sample an image sequence for you and send you the outcome free of any charge, to check EIKONA3D suitability for your work.
Fully operational JEOL JEM 100B with a W filament available in Export, = PA. Also includes numerous spare components. ALL offers considered, = best offer accepted.
For further information, contact:
Tom Isabell (724) 325-5444 tc_isabell-at-fischione.com
Thomas C. Isabell, Ph.D. Research Scientist E.A. Fischione Instruments, Inc. tc_isabell-at-fischione.com webpage: www.fischione.com
Hello all, Has anyone experienced puckering of tissues embedded in LRWhite resin? My = tissues are cell cultures fixed in 2%PFA, .5%GA, dehydrated to75% ETOH = then into graded LRWhite resin. I cut on a diamond knife and embed on 200 = mesh grids. Before staining, the puckers and wrinkles are seen as small, = and frequen, folds over most cells or along the cell borders. The resin = areas are totally free of folding. It's as if the tissue area is picking = up H2O during sectioning then has no where to go when it re-dries....very = frustrating. After staining it's worse, as the stain seems to stay in the = folds and gets really dark. Any thoughts and suggestions are very welcome = as always. Linda Fox Loyola University=20 Stritch School of Medicine lfox1-at-wpo.it.luc.edu
So called "osmium pepper" may be caused by the following:
1. Incompletely depolymerized paraformaldehyde or old glutaraldehyde that has started to polymerize. The aldehyde "globs" are left behind in the tissue to then react with the osmium and form a very fine precipitate. Do you see the pepper only in tissue? If so, this is a possibility.
2. Incomplete removal of PO4, as described by Richard Lander.
3. Lead stain that is precipitating (just starting to go bad). Is the pepper everywhere (even where tissue is absent)? If so, this is a possibility. Do unstained sections show the pepper? If not, then stain can be ruled out.
Best solutions: use freshly prepared, EM-grade aldehydes, cacodylate buffer, verified hi quality lead stain.
} } I am looking for a protocol for intestinal tissue. I have opossum } } intestinal tissue fixed by a colleague that has a lot of OsO4 pepper. I } } have also received gut tissue that appeared to have undergone autolysis. I } } am expecting more of these samples and would like to recommend a change in } } protocol. It is probably not possible to use perfusion.
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing 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 ####################################################################
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Manhattan, New York City:
If I could be helpful to your lab or imaging department, I may accept payment in exchange for education: as an intern, or barter, or as a volunteer "tryout". (Salary when appropriate.)
-------------
-Currently I work as a commercial web and print designer and use Photoshop imaging software daily for the past 4 years. -I have been freelancing; coloring microscans in photohop for a stock photography company. -Also, well practiced in nature photography -I have designed web pages* for the invertebrate scientists at the American Museum of Natural History and examined the light micoscope in the histology lab. I have also observed the Interdepartmental Labs' SEM and Confocal. As a layman, I find the visual aspect of scientific analysis is fascinating. *http://research.amnh.org/invertebrates *http://www.renorex.com/idltest (under construction)
Art and Science: I would like to continue combining my visual imaging skills with my general interest in Science. How can I learn microscopy equipment and/or 3D animated software and thereby provide assistance to scientists.
Thank you in advance for any education or vocation advice you have time to give.
Renee Recker 16 W. 16th St. NYC, NY 10011 212-675-1665
http://www.renorex.com (portions require shockwave and Netscape 4.0 browser)
P.S.
Previously I have posted e-mail asking about how to obtain microscopy education in a college program. I am still amazed and cannot thank enough the generous personal and professional responses. But, still stuck at the starting gate. The only course in commuting distance is NYU Spring '99. So, I thought I might ask about on-the-job training.
A few years ago I prepared some intestinal tissue for use in a probiotics study. Although my main aim was the fixation of the mucus blanket, it was also necessary to fix the intestine itself.
The protocol I used was an unconventional one - an anhydrous fixation, where osmium in solvent was used as the primary fixative. No "pepper" was observed, but the images, of course, are different from ones obtained using a conventional protocol where glutaraldehyde in buffer was used as the primary fixative. This may be OK for you, depending on what you want to see.
The following reference gives the details of the protocol, and shows the SEM results. The equivalent TEM results, which are as yet unpublished, looked good, too.
Allan-Wojtas, P., Farnworth, E.R., Modler, H.W. and Carbyn, S. (1997). A solvent-based fixative for electron microscopy to improve retention and visualization of the intestinal mucus blanket for probiotics studies. Microscopy Research and Technique 36:390-399.
If you would like a reprint, please contact me offline with your address and other necessary information, and I send one to you.
Just another idea.
Good Luck.
Paula.
Paula Allan-Wojtas Food Microstructure Specialist Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
I have been asked to post the following question. One of our students wants to selectively stain polystyrene (to differentiate from polyethylene oxide) and has read that this has been done by ruthenium oxide vapour staining. Has anyone had any experience with this that they could share with us?
TIA,
Pat Hales McGill University Dept. of Anatomy & Cell Biology hales-at-medcor.mcgill.ca
Yes I see it. There is apparently a server at okstate.edu which may be the culprit. I have added them as well as the 2 addresses which are associated to the bounding mail
Rich Lander and Y Chen
to the filter. That should stop the mail from reaching the rest of you until we can figure out why it started.
Richard and Ya you'lll be in libo for awhile. You should be able to receive Email from Microscopy but not post messages.
Dear Listers, I will need to reorder Mo grids. The (very old) ones I've been using have very ragged grid bars--often with holes in them--and I'd like to find some better-quality ones. Has anyone found a vendor for really good Mo grids? Any vendor who thinks (s)he qualifies is welcome to email me directly. TIA. Yours, Bill Tivol
Project MICRO has been contacted by an energetic 4th grade teacher in Marin County, CA. She's been doing some microscopy and wants to do more, using the new MSA/LHS "Microscopic Explorations". She needs help now with cleaning her scopes, and could use a volunteer to help with the microscopy later. Do any of you folks live in Marin? Reply offlist, to me or direct to her (cc to me, please?)
Caroline Schooley Educational Outreach 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/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/pci.html
I would like to contact Dr. O.P. Ottersen, who works in the department of Anatomy at the University of Oslo, Norway. Does anyone have his e-mail address, per chance?
Thank you in advance,
Doug ---------------------- Douglas R. Keene Associate Investigator Shriners Hospital Microscopy Unit 3101 S.W. Sam Jackson Park Road Portland, Oregon 97201 503-221-3434 DRK-at-shcc.org
The needle valves (vent, fill and drain) on our Ladd critical point drier are quite diffucult to turn, even when warm. Does anyone know if they can be lubricated and, if so, how?
Bob
Dr. Robert R. Wise Department of Biology and Microbiology University of Wisconsin-Oshkosh Oshkosh, WI 54901
(920) 424-3404 tel (920) 424-1101 fax wise-at-uwosh.edu www.uwosh.edu/departments/biology/wise/wise.html
I have a method for an Oil Red O stain for GMA sections. It uses 60% aqueous triethyl phosphate and 5% aqueous ferric ammonium sulphate, however I can only locate these chemicals in the powder form. Does anyone know if it matters if the powder form is used and what they are likely to be dissolved in as it doesn't state this in the methodology?
Elizabeth Cox Fisheries Biologist Queensland Department of Primary Industries Northern Fisheries Centre, PO Box 5396, Cairns Qld Australia 4870
Does anayone on this network have experience with the new field of Raman Imaging ? We have a raman instrument with a tunable filter and imaging. I have a couple of questions. I am a TEM person and new at this so excuse me if these questions are too elementary.
The manufacturer claims that on aquiring the image the spatial resolution of the 'spectra' is as small as a pixel. But since unlike IR, Raman is a scattering process wouldn't all regions limited by the beam (probe) , about 150 microns, be affected by each other and thus negating the claim of 1 pixel resolution? Infact I did observe this experimentaly while imaging composite polymeric samples. Spectra from all regions illuminated by the beam are almost identical although the image does exhibit contrast due to different phases.
Thanks
Kalpana
******************************************************************************** **** Kalpana S Katti, Ph.D. Department of Polymers and Coatings North Dakota State University Fargo, ND 58105 ph:(701)231-8410 fax:(701)231-8439 email:kkatti-at-prairie.nodak.edu ******************************************************************************** ****
} ------------------------------------------------------------------------ } 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, } Has anyone experienced puckering of tissues embedded in LRWhite resin? My tissues are cell cultures fixed in 2%PFA, .5%GA, dehydrated to75% ETOH then into graded LRWhite resin. I cut on a diamond knife and embed on 200 mesh grids. Before staining, the puckers and wrinkles are seen as small, and frequen, folds over most cells or along the cell borders. The resin areas are totally free of folding. It's as if the tissue area is picking up H2O during sectioning then has no where to go when it re-dries....very frustrating. After staining it's worse, as the stain seems to stay in the folds and gets really dark. Any thoughts and suggestions are very welcome as always. } Linda Fox } Loyola University } Stritch School of Medicine } lfox1-at-wpo.it.luc.edu } } Hi, You have several problems here. First, acrylics like LR White, etc., do not bond WITH the tissue as do epoxies. This allows a lot of shifting as soon as the stresses are relieved by cutting a thin section. LR White also is modestly poorly crosslinked (as compared to the epoxides). You can increase the crosslinkage chemically (unless you are doing immunostaining, then you do not want to heavily crosslink). What to do? Do you need to use LR White? What is your purpose of using it. It has a number of downsides which one can trade off in the immunoprocessing protocols for better location of antigens. For standard TEM work is is inferior to epoxides (wrinkling of thick sections, poor crosslinkage, polymerization damage, difficulty sectioning because of its property of attracting water to the block face, etc.) What to do? If you must use LR White, try using a 100 mesh grid with a film. Also when picking up floating grids, DO NOT suck the water off under the grid with filter paper. Do draw the water off by sticking filter paper between the forceps. Then, lay the grid in the forcepts down to dry. Pull a lamp with a 60W bulb down over the forceps to slightly warm the situation. Use at least 5 forceps at once. That way some grids are drying and others are being picked up. This is the recommendation of Newman who holds the patent for LR White and LR Gold (He sold the license to the London Resin Company.) Should you decide to more heavily crosslink the resin, contact EMS for advice and the chemical. (I have no stock in EMS). If your sections have folds and you posstain them with heavy metals, the stain will accumulate in the folds. There is nothing you can do about that. You must avoid the wrinkles in the first place. Should you have more trouble, contact me. Bye, Hildy
Pat: Both polystyrene and Polyethylene oxide will stain with ruthenium oxide. However they may stain at different rates. There is a paper by Trent et al in Macromolecules 1983 v16 pg. 589-598 that descibes RuO4 staining of polymers. RuO4 is quite hazardous so be aware of the safety issues. Steve
At 01:12 PM 9/10/98 -0400, you wrote: } } I have been asked to post the following question. One of our students wants } to selectively stain polystyrene (to differentiate from polyethylene oxide) } and has read that this has been done by ruthenium oxide vapour staining. } Has anyone had any experience with this that they could share with us? } } TIA, } } Pat Hales } McGill University } Dept. of Anatomy & Cell Biology } hales-at-medcor.mcgill.ca } } }
------------------------------ Stephen McCartney Research Associate Virginia Tech Materials Institute 2108 Hahn Hall Blacksburg, VA 24061-0344 USA
John Mike Taylor of the Electrical Engineering Dept at the University of Delaware is in need of price and delivery of a 40 pin EPROM made by Intel or compatible No. 8755AD or D8755A. If anyone out there has any source please contact John Mike Taylor directly at jtaylor-at-ee.udel.edu. Thanks in advance Pete Dondl, P & S Products
Responding to the message of {v04003a09b21dc32600cf-at-[141.233.130.134]} from "wise-at-vaxa.cis.uwosh.edu"-at-Sparc5.Microscopy.Com: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } The needle valves (vent, fill and drain) on our Ladd critical point drier } are quite diffucult to turn, even when warm. Does anyone know if they can } be lubricated and, if so, how?
I have the same problem with my Ladd CPD. Other than that, I find the Ladd CPD to be a very good device (IHNCIIL,JASC). I allow users of the CPD to loosen the valves with pliers, but NOT to tighten them that way, just use finger pressure to tighten. But I suspect that those with delicate fingers cheat a little when I'm out of the room, but hopefully, without over-tightening the valves. So far so good. We've had the unit for over 12 years and never had to replace a valve.
Is it possible to adjust the turning tension on these needle valves??
Gib Ahlstrand Electron Optical Facility, University of Minnesota, Dept. Plant Pathology 495 Borlaug Hall, St. Paul, MN. USA. 55108 (612)625-8249 612-625-9728 FAX, giba-at-puccini.crl.umn.edu
Michael T. Marshall Research Engineer, Electron Microscopy University of Illinois at Urbana-Champaign Frederick Seitz Materials Research Laboratory 104 South Goodwin avenue Urbana, IL 61801-2985 (217) 244-8193 fax: (217) 244-2278
Hi =20 Can anyone explain what the shape factor is (image analyses softwares) =20 As I understand it, when it is equal to 1 it means the object is a=20 disc...what if it is say 2.5 and 0.3?? =20 =20 Thanks =20 F.
wise-at-vaxa.cis.uwosh.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 } -----------------------------------------------------------------------.} } The needle valves (vent, fill and drain) on our Ladd critical point drier } are quite diffucult to turn, even when warm. Does anyone know if they can } be lubricated and, if so, how? } } Bob } } Dr. Robert R. Wise } Department of Biology and Microbiology } University of Wisconsin-Oshkosh } Oshkosh, WI 54901 } } (920) 424-3404 tel } (920) 424-1101 fax } wise-at-uwosh.edu } www.uwosh.edu/departments/biology/wise/wise.html
Dear Dr. Robert R. Wise,
RE: NEEDLE VALVES ON CPD
Try graphite or teflon grease. If this doesn't help than they may be wearing out and will probably need replacing if the problem gets too severe.
John Arnott --
LADD RESEARCH 13 Dorset Lane Williston, VT 05495
TEL 1-800-451-3406 (US) or 1-802-878-6711 (FROM ANYWHERE) fAX 1-802-878-8074 e-mail ladres-at-worldnet.att.net web site http://www.msa.microscopy.com/SM/LADD
} Does anayone on this network have experience with the new field of Raman } Imaging ? We have a raman instrument with a tunable filter and imaging. I } have a couple of questions. I am a TEM person and new at this so excuse } me if these questions are too elementary. } } The manufacturer claims that on aquiring the image the spatial resolution } of the 'spectra' is as small as a pixel. But since unlike IR, Raman is a } scattering process wouldn't all regions limited by the beam (probe) , } about 150 microns, be affected by each other and thus negating the claim } of 1 pixel resolution? Infact I did observe this experimentaly while } imaging composite polymeric samples. Spectra from all regions illuminated } by the beam are almost identical although the image } does exhibit contrast due to different phases.
I assume that you have an instrument wherein the sample is imaged onto a CCD, but in between there is a tunable filter so that at any one instant only a narrow range of wavelengths is striking the CCD. In this case light that is scattered from the sample will be imaged onto the CCD in the place corresponding to the place on the sample it came from.
I have been reading a lot of patents lately and it has made my exposition a little stilted, sorry.
Hope this helps. best regards mark
Mark W. Lund, PhD Director } } Soft X-ray Web page http://www.moxtek.com { { MOXTEK, Inc. 452 West 1260 North Orem UT 84057 801-225-0930 FAX 801-221-1121 lundm-at-xray.byu.edu
Jack and Jill go to court. Jill takes the stand and lies her head off. Jack takes the stand and lies his head off. Who's telling the truth?
We have an opening for a post-doctoral research associate to work on a project involving in situ straining of fcc metals. The purpose is to study dislocation intersections and to compare experimental observations with large-scale atomistic simulations. Experimental techniques include dynamic imaging, weak-beam dark-field microscopy and high resolution electron microscopy. Experience with in-situ straining is highly desirable but not essential.
Please send resumes and the names of three references to the address below or attach to an e-mail reply. The post-doctoral program at Los Alamos is described at http://stb.lanl.gov/postdoc/postdoc.html
Terence E. Mitchell Laboratory Fellow Center for Materials Science MS-K765 Los Alamos, NM 87545 voice mail: 505-667-0938 fax: 505-665-2992 e-mail: temitchell-at-lanl.gov http://www.mst.lanl.gov/cms/welcome.html http://www.mst.lanl.gov/CMS/EMF/emf-people.html
I need to get in touch with a company that used to be called GW Electronics and they used to be in Norcross, Ga. My "new address and phone number" card is dated Sept. 1, 1983. Are they still around?
Bill -- ============================================================= Bill Chissoe III Electron Microscopist,University of Oklahoma E-mail: wchiss-at-ou.edu Ph. (405)325-4391 =============================================================
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi,
Shape factors can be really useful for morphometry studies but I've seen some poor formulas used that miss a lot of detail. You have to look at the particular formula to see what the investigator is trying to describe, hopefully, they would give you a few examples of the object of interest and its value so the reader can gauge the meaning and significance of different values.
A good stereology text (unpaid endorsement) that talks a bit about this that I would recommend is:
Russ, J. C. (1986). Practical Stereology. New York, Plenum Press.
I think these journal articles might be helpful too:
True, L. D. (1996). "Morphometric applications in anatomic pathology [Review]." Human Pathology 27(5): 450-467.
Royet, J. P. (1991). "Stereology: a method for analyzing images." Progress in Neurobiology 37(5): 433-74.
Prakash, Y. S., K. G. Smithson, et al. (1993). "Measurements of motorneuron somal volumes using laser confocal microscopy: Comparisons with shape-based stereological estimations." Neuroimage 1: 95-107.
Lastly, a great resource (like this discussion list) for image analysis questions is to go to the NIH-Image website at http://rsb.info.nih.gov/nih-image/, and do a Gopher search of the discussion list archive, which is a terrific resource (plus join the discussion list!). If you didn't know, NIH-Image is free image analysis software, and has a bunch of free macro code that can be easily modified to fit many image analysis needs, and it is still FREE! Yahoo!
good hunting,
Brian C. Tryon MD/PhD student Allegheny University of the Health Sciences School of Medicine 3200 Henry Avenue Philadelphia, PA 19129 USA ----------------------------------------- "Quantifying is a committing task." - Cruz-Orive, 1994.
"For a successful technology, reality must take precedence over public relations, for Nature cannot be fooled." - Richard Feynman
} Can anyone explain what the shape factor is (image analyses softwares)
}
} As I understand it, when it is equal to 1 it means the object is a
} disc...what if it is say 2.5 and 0.3??
}
}
} Thanks
}
} F.
Hi,
Shape factors can be really useful for morphometry studies but I've seen some poor formulas used that miss a lot of detail. You have to look at the particular formula to see what the investigator is trying to describe, hopefully, they would give you a few examples of the object of interest and its value so the reader can gauge the meaning and significance of different values.
A good stereology text (unpaid endorsement) that talks a bit about this that I would recommend is:
{fontfamily} {param} Geneva {/param} {bigger} {bigger} Russ, J. C. (1986). {underline} Practical Stereology {/underline} . New York, Plenum Press.
{/bigger} {/bigger} {/fontfamily} I think these journal articles might be helpful too:
{fontfamily} {param} Geneva {/param} {bigger} {bigger} True, L. D. (1996). "Morphometric applications in anatomic pathology [Review]." {underline} Human Pathology {/underline} {bold} 27 {/bold} (5): 450-467.
Royet, J. P. (1991). "Stereology: a method for analyzing images." {underline} Progress in Neurobiology {/underline} {bold} 37 {/bold} (5): 433-74.
Prakash, Y. S., K. G. Smithson, et al. (1993). "Measurements of motorneuron somal volumes using laser confocal microscopy: Comparisons with shape-based stereological estimations." {underline} Neuroimage {/underline} {bold} 1 {/bold} : 95-107.
{/bigger} {/bigger} {/fontfamily} Lastly, a great resource (like this discussion list) for image analysis questions is to go to the NIH-Image website at http://rsb.info.nih.gov/nih-image/, and do a Gopher search of the discussion list archive, which is a terrific resource (plus join the discussion list!). If you didn't know, NIH-Image is free image analysis software, and has a bunch of free macro code that can be easily modified to fit many image analysis needs, and it is still FREE! Yahoo!
good hunting,
{fontfamily} {param} Geneva {/param} Brian C. Tryon
MD/PhD student
Allegheny University of the Health Sciences
School of Medicine
3200 Henry Avenue
Philadelphia, PA 19129
USA
-----------------------------------------
{/fontfamily} "Quantifying is a committing task." - Cruz-Orive, 1994.
"For a successful technology, reality must take precedence over public
relations, for Nature cannot be fooled." - Richard Feynman
At 03:13 PM 9/11/98 +0000, Frank S. wrote: } Can anyone explain what the shape factor is (image analyses softwares) } } As I understand it, when it is equal to 1 it means the object is a } disc...what if it is say 2.5 and 0.3??
It all depends on who wrote the code. Our old LeMont made a point of reporting what parameters were being used to calculate shape. Nowadays it is harder to tell.
Shape is often a ratio of width to length or vice versa. Whether the values are greater than 1 or less than 1 will give you a hint. Then there is the question of how W and L are calculated. They might be minimum and maximum projected measurements. L may be taken as the maximum projected measure and W as the projected measurement at 90 degrees to the L direction. They might be rectangular or elliptical equivalent measures given the features area and perimeter. Or they might be a ratio between perimeter squared and area. And that should, but may not always be, corrected by 4*pi.
Now if you are still using the Visilog package, the answer is Perimeter^2/(4*pi*Area). You should be able to plug in some actual measurements to verify it. A shape of 1 corresponds to a perfect circle - values larger than 1 indicate more elongation and/or convolution. You better not get any (many) values less than 1.
The shape factor of an object is defined by (4 X pi) X area / perimeter squared. In this instance a circle would have a shape factor of 1 and an irregularly shaped object would have a shape factor less than 1. Another way to measure shape is the area perimeter length squared / area. In this latter instance a circle would have a value of 4 x pi (12.57) and irregularly shaped objects would have values greater than this.
John
Can anyone explain what the shape factor is (image analyses softwares)
As I understand it, when it is equal to 1 it means the object is a disc...what if it is say 2.5 and 0.3??
Thanks
F.
John J. Turek, Ph.D. Associate Professor Department of Basic Medical Sciences 1246 Lynn Hall, G193C Purdue University W. Lafayette, IN 47907-1246 Phone: 765-494-5854 Fax: 765-494-0781 Email: jjt-at-vet.purdue.edu
The Chesapeake Society for Microscopy Fall Dinner Meeting
October 13th, 1998 Speaker: Dr. Joseph Gall (after dinner) "Microscopes and Discoveries in Cell Biology: Chicken or Egg?" 6:00 PM Social Hour 7:00 PM Dinner ($20.00, Students $10:00) Make a selection of one at time of reservation Maryland Crab Cakes Assorted Seafood Platter (fried) Filet Mignon with Mushrooms Roast Prime of Beef (thick cut) Pasta vegetarian dish Location: Snyders Willow Grove (410-789-8244) Linthicum, MD Make Reservation and meal selection by Oct 9th To: Andrea Weisberg-(301) 435-1977 aweisberg-at-nih.gov Dinner payable at meeting to 'CSM' Andrea S. Weisberg NIH/NIAID/LVD Bldg.4/Rm.210 4 Center Dr. Bethesda,MD 20892-0445 office (301) 435-1977 Fax (301) 480-1147 e-mail: aweisberg-at-nih.gov
I have an old Balzers CPD, circa 1979. It looks like someone has dropped the cover and chipped the glass window. I am concerned about its safety and need to do something to fix it.
This unit is pretty old and I need some help with repairs or parts. The glass window looks impossible to relace easily, looks like I need a whole new cover. It's a pretty substantial piece of stainless steel, threaded to go on the chamber, with a thick glass window pressed into it.
Does anyone have a lead on getting replacement parts or ideas about a safe repair?
If fixing is not possible, can anyone get me up to date on current CPD's to replace it?
Thanks.
Jonathan Krupp Microscopy and Imaging Lab University of California Santa Cruz, CA 95064 (831) 459-2477 FAX (831) 429-0146 jmkrupp-at-cats.ucsc.edu
I have a problem of cleaning components of EM like anodes, assemblies and aperatures when they are contaminated. Usual practice is to rub with diamond paste of various grades (depending upon contamination level) and ultrasonicating in solvent like methnol. Finally rinsing three-four times with fresh methonal. One SEM user has suggested me to ultrasonicate the items directly with liquid Ammonia and demonstrated it in his lab. Although it shows a very good cleaning, but I am fraid of whether they really works in EM.
As I have not come across any such reference can anybody experienced in this field, kindly guide me?
Thanks in advance.
Rajdeep Dongre Electron Microscopy Laboratory Agharkar Research Institute G.G. Agarkar Road, Pune - 411 004 India Phone : 91-212-353680/354357 91-212-351542 E-mail: rajdeep-at-aripune.ernet.in
The shape factor is defined as 4 * PI * area / (perimeter^2).
For a circle this computes to 1. Since a circle has the largest area for a given perimeter, any other geometrical figure will have a shape factor smaller than 1. Due to pixelation artifacts, sometimes values of larger than 1 are calculated, but these are artifacts that mostly appear for very small particles.
Hope that helps.
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St. # 105N Lakewood, CO 80215 voice: (888) FIND-SIS fax: (303) 234-9271 info-at-soft-imaging.com http://www.soft-imaging.com
} -----Original Message----- } ---------- } From: } "frank.sarrazit-at-AVESTASHEFFIELD.COM"-at-sparc5.microscopy.com[SMTP:"frank.sarra } zit-at-AVESTASHEFFIELD.COM"-at-sparc5.microscopy.com] } Sent: Friday, September 11, 1998 9:13 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: Shape factor (image analysis) } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} It all depends on who wrote the code. Our old LeMont made a point of } reporting what parameters were being used to calculate shape. Nowadays it is } harder to tell. snip } Now if you are still using the Visilog package, the answer is } Perimeter^2/(4*pi*Area). You should be able to plug in some actual } measurements to verify it. A shape of 1 corresponds to a perfect circle - } values larger than 1 indicate more elongation and/or convolution. You better } not get any (many) values less than 1.
In addition, we have used a similar 3-dimensional factor derived from Green (1927): sf= (36 x pi x vol-squared)/surface-area-squared for which sphere =1, cube=0.52 and a unit cylinder=0.67
Gib Ahlstrand wrote: } } } Responding to the message of {v04003a09b21dc32600cf-at-[141.233.130.134]} } from "wise-at-vaxa.cis.uwosh.edu"-at-Sparc5.Microscopy.Com: } }
} } The needle valves (vent, fill and drain) on our Ladd critical point drier } } are quite diffucult to turn, even when warm. Does anyone know if they can } } be lubricated and, if so, how? } } I have the same problem with my Ladd CPD. Other than that, I find the Ladd CPD } to be a very good device (IHNCIIL,JASC). I allow users of the CPD to loosen the } valves with pliers, but NOT to tighten them that way, just use finger pressure } to tighten. But I suspect that those with delicate fingers cheat a little when } I'm out of the room, but hopefully, without over-tightening the valves. So far } so good. We've had the unit for over 12 years and never had to replace a valve. } } Is it possible to adjust the turning tension on these needle valves?? } } Gib Ahlstrand
Dear Gib Ahlstrand,
Thank you for your kind comments about the Ladd CPD. You can adjust the tension nut next to the foam behind the needle valves. A word of caution though, you must be careful not to overloosen as it may start to leak. You can also remove the tension nut, take the stem from the valve and apply graphite, white lithium or teflon grease to make the needle valves easier to turn. Too much grease may clog the valve so it should be applied conservativly. If you have any further questions you would like to discuss, please feel free to call Mike Bouchard here at Ladd at 1-802-878-6711. Hope this is of some help,
John Arnott Chairman --
LADD RESEARCH 13 Dorset Lane Williston, VT 05495
TEL 1-800-451-3406 (US) or 1-802-878-6711 (FROM ANYWHERE) fAX 1-802-878-8074 e-mail ladres-at-worldnet.att.net web site http://www.msa.microscopy.com/SM/LADD
I've done cell culture using LRWhite fairly similarly to how you treated your cells. I dehydrated to 100% ethanol, which may have helped with infiltration. I didn't do it, and I don't know if you did, but embedding in a vacuum might help.
When was working on these cells, the only dishes the cells would grow on were slightly soluble in LRWhite. Where the plastic dissolved, it was opaque white, covering the cells. I asked the company if there was a way around that, they said that it shouldn't interfere with anything I was doing. So I sectioned through the dish and the LRWhite, and was able to immunostain. I don't know to what degree, but I guess I embedded my cells in LRWhite and the soluble component of tissue culture plastic. Maybe that stiffened it to prevent the folding problem you're having. Hope this is helpful. Charlie Ginsburg Research Dept. National College of Chiropractic Lombard IL
Linda Fox {lfox1-at-wpo.it.luc.edu} wrote: } } Hello all, } Has anyone experienced puckering of tissues embedded in LRWhite resin? My tissues are cell cultures fixed in 2%PFA, .5%GA, dehydrated to75% ETOH then into graded LRWhite resin. I cut on a diamond knife and embed on 200 mesh grids. Before staining, the puckers and wrinkles are seen as small, and frequen, folds over most cells or along the cell borders. The resin areas are totally free of folding. It's as if the tissue area is picking up H2O during sectioning then has no where to go when it re-dries....very frustrating. After staining it's worse, as the stain seems to stay in the folds and gets really dark. Any thoughts and suggestions are very welcome as always. } Linda Fox } Loyola University } Stritch School of Medicine } lfox1-at-wpo.it.luc.edu }
_________________________________________________________ DO YOU YAHOO!? Get your free -at-yahoo.com address at http://mail.yahoo.com
Contamination removal has become a very hot topic and has prompted a lot = of research into methods to remove it. We have commercialized a plasma cleaning system based on technology developed by Dr. Nestor Zaluzec (our friendly neighborhood sysop!). We have collaborated on a lot of research=
into plasma cleaning for electron microscopy and have done a fair amount = of work on cleaning microscope parts and accessories. =
While our system was actually developed for cleaning TEM specimens and specimen holders, it can also be used conveniently for cleaning any type = of microscope parts. The only limitation is that it must be able to fit within the 6" ID and 4" tall chamber - not a problem for almost anything you would like to clean. Please contact me off-line or visit our web sit= e for more detailed information.
Elisabeth: Aqueous means water based. So you need to make a 60% and a 5% solution of those chemicals in water. A percent solution of a solid in water solution means somany grams made up to 100 ml. So its 60 grams made up to 100ml when fully dissolved - not many powders will dissolve in that concentration. Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au
On Friday, 11 September 1998 11:37, Cox, Elizabeth [SMTP:CoxE-at-prose.dpi.qld.gov.au] wrote: } I have a method for an Oil Red O stain for GMA sections. } It uses 60% } aqueous triethyl phosphate and 5% aqueous ferric ammonium } sulphate, however } I can only locate these chemicals in the powder form. } Does anyone know if } it matters if the powder form is used and what they are } likely to be } dissolved in as it doesn't state this in the methodology?
} Elizabeth Cox } Fisheries Biologist } Queensland Department of Primary Industries } Northern Fisheries Centre, } PO Box 5396, } Cairns Qld Australia 4870 } } ph:+61 7 4035 0100 } Fax: +61 7 4035 1401 } } }
Sally, about 15 years ago somebody published the reason for this Os "pepper". Simply, for this to occur, some chemically unbound Os, GA and phosphate must remain in the tissue. If any one of these is missing you will not get this frustrating artefact.
I suppose this is one major reason for the popularity of cacodylate buffer. It makes the very thorough rinsing process which is required between GA and OS redundant; simply any unbound GA no longer matters.
The situation is aggravated by your fixation schedule. That tissue is overfixed! One hour in 1% GA and one hour in one percent OSO4 in the cold is the general standard for fixing of soft tissues. Intestine should be easy and which animal the tissue is from would not matter for fixation purposes. Fixing at room temperature may be two to four times more severe than fixing "on ice". Your fixation schedule should be close to the point were the osmium completely oxidised membranes and "white" spaces only remain. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au *****
On Thursday, 10 September 1998 6:13, Sally Burns [SMTP:burnssal-at-pilot.msu.edu] wrote: } Gut tissue fixation } I am looking for a protocol for intestinal tissue. I have } opossum } intestinal tissue fixed by a colleague that has a lot of } OsO4 pepper. I } have also received gut tissue that appeared to have } undergone autolysis. I } am expecting more of these samples and would like to } recommend a change in } protocol. It is probably not possible to use perfusion. } I am looking for intercellular and intracellular bacteria } and /or } protozoal infections in these tissues. } Would the chemistry of the intestine be causing this type } of problem? Or } is it simply a matter of inadequate washing? } Thanks??.. Sally } } The fixation used was: } 2% glut in PBS at pH 7.2 for 3 hours. } 3X washed in PBS } 1X washed in H20 } 1% OsO4 for 4H room T } washed 3x in H20 } dehydrated in a 25% series of acetone } Infiltrated and embedded in a 25% series into a Spurrs/ } Quetol blend. } } } } Sally Burns } Center for Electron Optics } B5 Pesticide Research Center } (517) 355-5004 } } burnssal-at-pilot.msu.edu }
In theory, I expect that when compared with a tungsten filament, at least the solid, single post LaB6 cathode design by Kimball, would make for less movement, especially during the warming-up period. In practise, tungsten filaments are more stable because the emitting area is much larger and so, minor misalignment due to a drifting filament is less consequential. The stability we are talking about is due to slow drift and this is of no consequence to normal, including high resolution imaging in TEM or SEM. Stability over several minutes matters when performing quantitative microanalyses with a probe or EDS in dedicated SEM/TEM. In quantitative analyses a spectrum maybe acquired over two minutes and that is related to standard spectra and all numbers must lead to results that come within 1% of reality. The obvious way to increase stability in LaB6 is to increase the size of the microflat at the tip of the LaB6 cone. The normal size of the cone (Kimball's) is 15 or 20 micrometer. A 40 micrometer microflat is also available (unfortunately at greater cost) and this makes a LaB6 quite suitable for quantitative microanalyses. Brightness is reduced but for microanalyses that is no consideration. Reducing downtime by increasing "filament life" to over 5000 hours at low and medium emission is the main benefit of a Lab6 in a microprobe. For non-analyses work, a standard flat Lab6 cathode is quite stable for SEM and TEM requirements; extreme drift would matter but stability worries in these applications are more likely to relate to HV instabilities or a pulsing beam due to column/aperture contaminations. The question for Mark Darus is the EM's vacuum system: Is it good enough for LaB6 operation. Disclaimer: ProSciTech supplies Kimball cathodes and filaments. Jim Darley
ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au *****
On Wednesday, 9 September 1998 0:38, Mark Darus [SMTP:DARUSM-at-cle.lg.bfg.com] wrote: } I'm new at operating an SEM that has a tungsten filament. } For the past 5 } years I had an instrument that used a LaB6, but I've } changed companies. } I have 2 questions. } 1. Both the people here and the instrument's service man } tell me } that the tungsten filament is more stable than the LaB6. } I ask them to } explain further and they really don't get into it. What } is the stable thing } about the tungsten filament? My LaB6 seemed fine to me, } perfect in fact, } so what is the unstableness of it? } 2. I'm also told here that, with the tungsten filament, } as the beam sits in } one area on the sample, Carbon will develop in that area. } Is this true? If } so how long does it take for the carbon to contaminate the } area, and also } does this take any confidence in a carbon analysis and } throw it out the } window? }
Just a reminder to all that, one of the good places to look for things like this is the SUSTAINING MEMBERS WWW page of MSA
http://www.msa.microscopy.com/SM/SustMembers.html
from that page the Current GW address is:
GW Electronics, Inc. Attn: Mr. Larry H. Glassman 6981 Peachtree Industrial Blvd. Norcross, GA 30092 Phone(days): (404) 449-0707 Fax Number: (404) 449-0284 E-Mail:
} } I need to get in touch with a company that used to be called GW } Electronics and they used to be in Norcross, Ga. My "new address and } phone number" card is dated Sept. 1, 1983. Are they still around? } } Bill } -- } ============================================================= } Bill Chissoe III } Electron Microscopist,University of Oklahoma } E-mail: wchiss-at-ou.edu Ph. (405)325-4391 } =============================================================
At 03:25 PM 9/3/98 +0900, =C1=A4=C1=F6=C8=C6 wrote:=20
} } } }
{excerpt} {fontfamily} {param} =B1=BC=B8=B2 {/param} {smaller} I am a freshman stu= dying dentistry in Seoul, Korea. Yesterday my professor gave us a question which we were supposed to answer via e-mail as quick as we could. The problem was "Why is there the 'phase contrast' in PCM?" I looked up all the references available to me, and searched all over the internet for 5 hours, and still don't have the answer. If can anybody help me, please e-mail me at { {mailto:gehoon-at-plaza1.snu.ac.kr} gehoon-at-plaza1.snu.ac.kr
{/smaller} {/fontfamily}
{/excerpt} { { { { { { { {
Hi,
The answer is yes, there very definitely is phase contrast in PCM. In regular brightfield microscopy, the image is formed by interference between the undiffracted background light and the light diffracted by the features in the specimen. This interference occurs at the Primary Image Plane, which can be easily viewed by removing the eyepiece and stretching a piece of lens paper over the resulting opening. The problem in viewing many unstained biological samples is that the phase relationship between the undiffracted background light and the diffracted specimen light is on the order of a quarter of a wavelength or less. The result is incomplete interference and very low contrast.
For Phase Contrast Microscopy, we carefully engineer the microscope to take advantage of this quarter wavelength phase shift. The whole concept is based on the the principle that, when waves are HALF a wavelength out of step, they will undergo destructive interference. Our challenge: to increase the phase shift between the background light and the specimen light to meet this requirement. The implementation is elegantly simple (so elegant, it earned Frits Zernike the Nobel prize!):
a. First, we need to control the exact location of the background light.=20 To accomplish this feat, we limit the aperture in the front focal plane of the condenser to just a ring.
b. Next, we need to generate the extra quarter wave difference. We accomplish this feat by inserting a special phase plate in the back focal plane of the objective. You can view this plane by rotating a phase objective in place then removing the eyepiece and looking far down the tube, into the back of the objective. There you will find a dark or "smokey" ring. =20
This back focal plane is "conjugate" (optically related) to the front focal plane of the condenser. If you rotate the phase annulus in the condenser into position, you will see the bright ring underlaying the smokey one. When the phase system is correctly aligned, you will notice that the bright ring from the condenser is imaged precisely in the darker ring mounted in the objective.=20
The phase plate which is mounted in the back focal plane of the objective has two characteristics:
One is a {underline} channel {/underline} , usually of less thickness than the general area of the plate. In most conventional phase systems, this channel is cut into the ring so that the background light has less glass to go through. The amount of the cut allows the background light to gain the extra quarter wave jump on the diffracted light from the sample.
The second characteristic of the phase plate is a {underline} neutral density filter {/underline} (the reason that this ring looks dark). For optimum interference, the two interfering waves need to be approximately the same amplitude. However, the diffracted light is usually only about 15% as bright as the background light. To correct this mismatch, we coat the cut through which the background light passes with a neutral density material.
The final outcome: when the diffracted light from the specimen meets the undiffracted light from the background in the Primary Image Plane, they are out of step by a total of one-half wave and are about the same intensity. They destructively interfere, generating darker features against a lighter (soft gray) background. Voila! Phase contrast.
c. One more point: To make the system work really well, you need to take into account several variables.
One is the sample: it really needs to be the type which closely approximates the initial quarter wavelength phase shift. Since this shift depends on both the real, geometric thickness of the specimen as well as the difference=20
in refractive index between the mounting medium and the sample, you can fine tune the system by changing the mounting medium (try glycerin, white corn syrup, or even immersion oil, if you sample will tolerate=20 it)
The second is the wavelength of light. Notice that we did not mention a quarter of a wave shift for any particular wavelength. Most modern phase contrast microscopes are engineered for 546 nm, so you need to use a good 546 nm (rich green) filter.
Finally, alignment is critical. Make sure that you have the correct phase ring or annulus in the condenser which matches the phase plate in the objective.
There are several really good references on Phase Contrast:
1. Ross, K. F. A. Phase Contrase and Interference Microscopy for Cell Biologists. Edward Arnold, Ltd, London. 1967
2. Pluta, M. Advanced Light Microscopy, Vol. 2.,Elsevier, NY. 1988
For a thorough but practical discussion, we also recommend our book, Optimizing Light Microscopy for Biological and Clinica Laboratories (1997). Details are available on our web site: { {http://www.MME-Microscopy.com/education}
Augusto, please check Micron 27(2) 1997, 129-139 and J. of Microscopy 188, 1997, 285-289.
-- Jaap Brink, Ph.D. Biochemistry, One Baylor Plaza, Baylor College of Medicine, Houston, TX 77030 Phone: (713)798-6989 -- Fax: (713)796-9438 -- Email: jbrink-at-bcm.tmc.edu URL : http://ncmi.bioch.bcm.tmc.edu/~brink
On Fri, 11 Sep 1998 Augusto_A_Morrone-at-notes.seagate.com-at-sparc5.microscopy.com wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } Subject: TEM imaging, CCD Vs film } } There have been several threads in the past on the advantages and } disadvantages of film and CCD cameras to acquire images in the TEM. I } enjoy having both capabilities as complementary in the operation of the } TEM, and became dependent on both. For convenience, I still take images } on film and then digitize the negatives; sometimes I also make contact } prints or 5x to 9x enlargements in the darkroom. However, I have no hard } data (numbers) to back up my impression that the resolution and contrast } range of film is better. Can anyone send me a note on this issue, a } reference, or forward me an earlier posting (couldn't find it in the } archives for the last several months)? } } Thank you. } } Augusto A. Morrone } Seagate Technology } 7801 Computer Ave South } Bloomington, MN 55435-5489 } Phone: (612) 844-5838 } Fax: (612) 844-8247 } Augusto_ A_Morrone-at-notes.seagate.com } } }
Augusto, please check Micron 27(2) 1997, 129-139 and J. of Microscopy 188, 1997, 285-289.
-- Jaap Brink, Ph.D. Biochemistry, One Baylor Plaza, Baylor College of Medicine, Houston, TX 77030 Phone: (713)798-6989 -- Fax: (713)796-9438 -- Email: jbrink-at-bcm.tmc.edu URL : http://ncmi.bioch.bcm.tmc.edu/~brink
On Fri, 11 Sep 1998 Augusto_A_Morrone-at-notes.seagate.com-at-sparc5.microscopy.com wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } Subject: TEM imaging, CCD Vs film } } There have been several threads in the past on the advantages and } disadvantages of film and CCD cameras to acquire images in the TEM. I } enjoy having both capabilities as complementary in the operation of the } TEM, and became dependent on both. For convenience, I still take images } on film and then digitize the negatives; sometimes I also make contact } prints or 5x to 9x enlargements in the darkroom. However, I have no hard } data (numbers) to back up my impression that the resolution and contrast } range of film is better. Can anyone send me a note on this issue, a } reference, or forward me an earlier posting (couldn't find it in the } archives for the last several months)? } } Thank you. } } Augusto A. Morrone } Seagate Technology } 7801 Computer Ave South } Bloomington, MN 55435-5489 } Phone: (612) 844-5838 } Fax: (612) 844-8247 } Augusto_ A_Morrone-at-notes.seagate.com } } }
Augusto, please check Micron 27(2) 1997, 129-139 and J. of Microscopy 188, 1997, 285-289.
-- Jaap Brink, Ph.D. Biochemistry, One Baylor Plaza, Baylor College of Medicine, Houston, TX 77030 Phone: (713)798-6989 -- Fax: (713)796-9438 -- Email: jbrink-at-bcm.tmc.edu URL : http://ncmi.bioch.bcm.tmc.edu/~brink
On Fri, 11 Sep 1998 Augusto_A_Morrone-at-notes.seagate.com-at-sparc5.microscopy.com wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } } Subject: TEM imaging, CCD Vs film } } There have been several threads in the past on the advantages and } disadvantages of film and CCD cameras to acquire images in the TEM. I } enjoy having both capabilities as complementary in the operation of the } TEM, and became dependent on both. For convenience, I still take images } on film and then digitize the negatives; sometimes I also make contact } prints or 5x to 9x enlargements in the darkroom. However, I have no hard } data (numbers) to back up my impression that the resolution and contrast } range of film is better. Can anyone send me a note on this issue, a } reference, or forward me an earlier posting (couldn't find it in the } archives for the last several months)? } } Thank you. } } Augusto A. Morrone } Seagate Technology } 7801 Computer Ave South } Bloomington, MN 55435-5489 } Phone: (612) 844-5838 } Fax: (612) 844-8247 } Augusto_ A_Morrone-at-notes.seagate.com } } }
Bill Chissoe III wrote: ================================================== I need to get in touch with a company that used to be called GW Electronics and they used to be in Norcross, Ga. My "new address and phone number" card is dated Sept. 1, 1983. Are they still around? ================================================== G-W Electronics is very much "still around". Their website is at URL http://www.gwelectronics.com/
and their address/contact information is the following:
=================================================== 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 ############################ ==================================================
Does anyone know of a source for an old Spencer-style external lamp with collector lens, diaphragm, filter holder, and bulb centering adjustment? Thanks.
-----Original Message----- } From: System Administrator To: Howe L. C. Josephine Sent: 9/13/98 11:24:59 AM
} ---------- } From: Howe L. C. Josephine } Sent: Sunday, September 13, 1998 11:24 AM } To: 'microscopy-at-msa. microscopy.com' } Subject: radiation from uranyl acetate } } A postgraduate had prepared 3 tubes of 10g uranyl acetate in 15 mls water. } Later she decided to dispose it away. She brought it to the radiactive } waste disposal to discard it. The safety officer did a check to see } whether it was safe to dispose it there. The radiation emitted was very } high, about 500 counts per unit. She was told it was not safe for her to } handle it without protection. She was worried and went for a blood test. } It showed the cell count of lymphocyte was lower than normal. 6 weeks } later she went for another count. This time the cell count was much } higher. She is now very worried and would like to how harmful is the } radiation from uranyl acetate. Can anybody help to ease her anxiety? } Till today she has not forgiven people who has been handling uranyl } acetate for not informing her of the risk. } } Josephine Howe } NUS }
} Does anayone on this network have experience with the new field of Raman } Imaging ? We have a raman instrument with a tunable filter and imaging. I } have a couple of questions. I am a TEM person and new at this so excuse } me if these questions are too elementary. } } The manufacturer claims that on aquiring the image the spatial resolution } of the 'spectra' is as small as a pixel. But since unlike IR, Raman is a } scattering process wouldn't all regions limited by the beam (probe) , } about 150 microns, be affected by each other and thus negating the claim } of 1 pixel resolution? Infact I did observe this experimentaly while } imaging composite polymeric samples. Spectra from all regions illuminated } by the beam are almost identical although the image } does exhibit contrast due to different phases.
This question can be best answered by the manufacturer of your instrument. However, I guess that your instrument has been designed to get the best possible resolution. In that case, as in other optical microscopy, the resolution is limited by the diffraction of light. In other words, due to the wave nature of light you can not focus a light beam below a certain size. In the visible range that means that your spatial resolution is around one micrometer on the sample.
Probably the manufacturer has arranged things in such a way that one micron on the sample is imaged at least as one pixel on the CCD detector.
I hope this helps.
--------------------------------------------------------------------------- Fernando Agullo'-Rueda Raman Microscopy Laboratory Instituto de Ciencia de Materiales de Madrid (CSIC) Cantoblanco, E-28049 Madrid Espan~a (Spain)
We normally use 'Quadralene' which is a detergent containing ammonia (followed by rinsing), although I have used ammonia when cleaning gun parts which aren't easy to remove from the gun. The only thing to remember is that you must never use ammonia on copper or brass parts (if you have any in your gun). Also I am surprised that you use methanol for cleaning - do you not think that ethanol or acetone are safer?
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD UK
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk ---------- } From: Rajdeep Dongre To: microscopy
Dear Friends,
I have a problem of cleaning components of EM like anodes, assemblies and aperatures when they are contaminated. Usual practice is to rub with diamond paste of various grades (depending upon contamination level) and ultrasonicating in solvent like methnol. Finally rinsing three-four times with fresh methonal. One SEM user has suggested me to ultrasonicate the items directly with liquid Ammonia and demonstrated it in his lab. Although it shows a very good cleaning, but I am fraid of whether they really works in EM.
As I have not come across any such reference can anybody experienced in this field, kindly guide me?
Thanks in advance.
Rajdeep Dongre Electron Microscopy Laboratory Agharkar Research Institute G.G. Agarkar Road, Pune - 411 004 India Phone : 91-212-353680/354357 91-212-351542 E-mail: rajdeep-at-aripune.ernet.in
id AA01120; Mon, 14 Sep 1998 10:12:55 -0400 Received: from FETC-Message_Server by FETC.DOE.GOV with Novell_GroupWise; Mon, 14 Sep 1998 10:16:27 -0400 Message-Id: {s5fcecfb.080-at-FETC.DOE.GOV} X-Mailer: Novell GroupWise 4.1
I have a way of doing this that I use regularly with good results in my study coal ash deposits---similar to rocks and fracture surfaces in ceramics. I use reflected-light illumination (bright- and dark- field) . The method can be easily implemented with typical image processing software---I have Optimas macros available. For more information: ftp://titan.petc.doe.gov/pub/ramer (Under /docs is a Word document of my Microscopy Today article [Feb/March 98], a PowerPoint file illustrating results, and an AVI file of an animated fly-by of a dot on the back of a penny. The macros are under /Optimas.)
Briefly, the method is: 1. Get a stack of images---The raised dot on a coin makes a good object: I use 5 um steps with a 20X objective and 2 um for 50X, which can be done easily by hand. 2. Save the focused part of each image---Apply the LAPLACIAN operator to find texture (scratches). Convert the result to a binary image (choosing the threshold will require some trial and error, but the threshold will remain the same for all the images in the stack). The binary image will have "snowy" patches corresponding to the regions with texture. Merge the "snow flakes" into solid patches by DILATING several times. AND the result with the original image and you should have the focused part. 3. Combine the focused parts of all the images---Use the MAX operator instead of ADD (or OR) because the focused parts might overlap some.
Everett Ramer Federal Energy Technology Center Pittsburgh, PA (412)892-4920
The ChemIcon instrument uses the LCTF as a tunable bandpass filter. The idea behind the instrument is to globally illuminate the sample with a large laser spot and then collect all of the Raman scatter from the sample and filter it through the LCTF.
You then can collect spectra by taking a series of images with the CCD, each at a different wavelength. If you select the same pixel in each of the wavelength dependent images and plot it's intensity as a function of the image wavelength, you can reproduce a spectra. Therefore, if you have a 100 pixel by 100 pixel image, you can have 10000 spectra in one spectral image dataset. The spectral resolution is based on the bandpass of the filter, which is about 7 cm-1. The spatial resolution, as Fernando said, is diffraction limited.
Although I no longer work for the compant, I helped design the ChemIcon instrument. So, if you have any questions about it's use or the science behind it's design, please feel free to contact me.
Tim Prusnick, PRUSNICK_TIM-at-worldnet.att.net Raman Application & Support Engineer Renishaw Inc. 623 Cooper Court Schaumburg, IL 60173 PHONE: 847-843-3666 FAX: 847-843-1744
-----Original Message----- } From: Fernando Agullo-Rueda [mailto:FAR-at-icmm.csic.es] Sent: Monday, September 14, 1998 2:35 AM To: microscopy-at-Sparc5.Microscopy.Com
Kalpana S Katti wrote:
} Does anayone on this network have experience with the new field of Raman } Imaging ? We have a raman instrument with a tunable filter and imaging. I } have a couple of questions. I am a TEM person and new at this so excuse } me if these questions are too elementary. } } The manufacturer claims that on aquiring the image the spatial resolution } of the 'spectra' is as small as a pixel. But since unlike IR, Raman is a } scattering process wouldn't all regions limited by the beam (probe) , } about 150 microns, be affected by each other and thus negating the claim } of 1 pixel resolution? Infact I did observe this experimentaly while } imaging composite polymeric samples. Spectra from all regions illuminated } by the beam are almost identical although the image } does exhibit contrast due to different phases.
This question can be best answered by the manufacturer of your instrument. However, I guess that your instrument has been designed to get the best possible resolution. In that case, as in other optical microscopy, the resolution is limited by the diffraction of light. In other words, due to the wave nature of light you can not focus a light beam below a certain size. In the visible range that means that your spatial resolution is around one micrometer on the sample.
Probably the manufacturer has arranged things in such a way that one micron on the sample is imaged at least as one pixel on the CCD detector.
I hope this helps.
--------------------------------------------------------------------------- Fernando Agullo'-Rueda Raman Microscopy Laboratory Instituto de Ciencia de Materiales de Madrid (CSIC) Cantoblanco, E-28049 Madrid Espan~a (Spain)
Dear Rajdeep, } } I have a problem of cleaning components of EM like anodes, assemblies } and aperatures when they are contaminated. Usual practice is to rub with } diamond paste of various grades (depending upon contamination level) and } ultrasonicating in solvent like methnol. Finally rinsing three-four times } with fresh methonal. One SEM user has suggested me to ultrasonicate the } items directly with liquid Ammonia and demonstrated it in his lab. } Although it shows a very good cleaning, but I am fraid of whether they } really works in EM. } We remove some of the harder-to-clean contamination with a polishing compound (similar to your diamond paste method), then we soak the parts-- mostly aluminum liners, stainless steel Wehnelt cylinders, and a few of other materials--in Alconox, a mild detergent, rinse with distilled water, rinse with ultra-pure water, rinse with ethanol (better for fingerprints & less toxic than methanol), and rinse with acetone. We do not sonicate; it is not a safe procedure with organic solvents, but most people get away with it. The microscopy list archives have considerable information about this. Good luck. Yours, Bill Tivol
Ruthenium tetroxide is a choice for polystyrene staining and since it is a strong oxidizing agent you will need to do a 1, 2, 3....min stainig time to determine the best time period for your sample. Be aware that the staining needs to be done under the hood-follow safety procedures- If you have any specific questions email me.
Ani
Ani M Issaian California Institute of Technology Pasadena, CA. 91125 MC 210-41
I am interested in high resolution coaters to use with field emission SEM. Does anyone working with FE-SEM's have some opinions for me on what is best specifically for polymer applications. Steve
------------------------------ Stephen McCartney Research Associate Virginia Tech Materials Institute 2108 Hahn Hall Blacksburg, VA 24061-0344 USA
Anyone doing yeast embedding for TEM immunocytochemistry on a routine basis, please contact me directly to discuss a possible collaborative work. Thanks in advance.
Michel **************************************************** Michel Deschuyteneer, Ph.D. deschuyt-at-sbbio.be Scientist Electron Microscopy Laboratory
SmithKline Beecham Biologicals Rue de l'Institut, 89 B1330 Rixensart, BELGIUM Tel: +32-2-656 9290 Fax: +32-2-656 8164 **************************************************** Standard disclaimer: the opinions expressed in this communication are my own and do not necessarily reflect those of SmithKline Beecham. ****************************************************
As most of the people on the web know Protrain have a history in EM servi= ce and operator training and that we regulary run courses on the maintenance=
of electron microscopes. =
A little plug for the University of Sydney, Australian Centre for Microscopy and Analysis, who will be hosting our "Monitoring & Maintainin= g the Electron Microscope" course from 26th to 29th October 1998. Contact = - emma-at-emu.usyd.edu.au - for details.
That said attached is a cleaning programme for electron microscopes that = I hope will help you out.
Steve Chapman
Senior Consultant E.M. Protrain, 16 Hedgerley, Chinnor, Oxford OX9 4TN, England. Tel & Fax 44 (0)1844 353161 Web Site - http://ourworld.compuserve.com/homepages/protrain For Consultancy and Courses in Electron Microscopy World Wide
Maintaining a Scanning Electron Microscope=0D =0D TUNGSTEN Gun Systems=0D =0D The cathode assembly should be cleaned every filament change, the anode e= very other change and the electron gun at least once a year.=0D =0D Materials - Almost any metal polish may be used to clean electron gun com= ponents however it must not be LONG LIFE. Long life additives coat the c= leaned item with a polymer that causes chaos in the electron gun. Look o= ut for any indication on the bottle or tube that the manufacturer is clai= ming that you will not need to clean the metalwork so often after using t= heir product!=0D =0D Method - Almost more important than the cleaning efficiency is our abilit= y to completely remove the polishing media. So many service call outs ar= e due to problems caused through inefficient removal of the media. For t= his reason it makes sense to use a metal polish that is easily removed by= a solvent for tungsten. In this way we not only remove the metal polish= but also clean the areas that are difficult to approach with the polish,= nooks and crannies! Also very important is the need to clean without da= maging the component, scratching it or placing cotton hairs within the "t= raps" that the manufacturers seem to put in our way. The best cleaning t= echnique is a wet clean, which is to use solutions and an ultrasonic clea= ner. In this way the damage that mechanical forces apply to the componen= ts are minimised. Sure the cathode aperture may need a little more encou= ragement to give up its deposit but only do this if the wet cleaning proc= edure falls short. We like "Silvo" or "Bluebell" or "Brasso", liquid met= al polishes that will mix with a dilute ammonia solution to form a cleani= ng media, but a solution that may be removed with further washes in dilut= e ammonia. The mix - 10% metal polish in 90% ammonia solution - where th= e solution is 10% ammonium hydroxide in water. Place the components, one= at a time, in the solution with their least important face down wards. = Never put gun components together in the solution, as they will damage ea= ch other. Do not put an aluminium cathode in ammonia as it will go black= , oxide! After 20 minutes in an ultrasonic the component should be clean= , wash off in running water and run for another 5 minutes in straight 10%= ammonium hydroxide in water. Swill off with running water and then wash= in alcohol and dry. NEVER throw away your solutions until you have reas= sembled the cathode, as it is quite possible for the small screws to have= fallen out and to reside in the debris at the base of one of the cleanin= g containers. If you do have a deposit remaining in the aperture area of= the cathode a little mechanical effort with the cleaning media may be re= quired.=0D =0D Once a component is clean check it with a hand lens or binocular microsco= pe before returning it to the microscope OR wrap it in kitchen (aluminium= ) foil until required. When working with clean components work on a shee= t of this foil as it is very clean and it makes an ideal working surface.= =0D =0D When rebuilding the components and placing them in the microscope try to = place the higher components first so that you are less likely to drop deb= ris on the cleaned components below. Alternatively always cover the colu= mn with aluminium foil when it is opened for removing components or close= the gun chamber down whilst cleaning is being carried out.=0D =0D The gun chamber IS important and this should be cleaned through disassemb= ly once a year, particularly with a TEM. Dirty guns hold gas and induce = micro discharge, which spoils images. Clean the gun chamber with metal p= olish, remove the metal polish with dilute ammonia and buff up the walls = with a clean chamois or dear skin leather. To retain the cleanliness of = the chamber, each time you change a filament buff up the walls with the l= eather. If the chamber smells, oily-ozone smell, but is not visibly stai= ned, this is the result of discharge and all traces of the smell should b= e removed with dilute ammonia.=0D =0D Look after your gun, it is probably the dirtiest area of the microscope, = other than the specimen area in a SEM or the camera chamber in a TEM, its= state will determine the ultimate performance of the instrument and your= filament life.=0D =0D LANTHANOM HEXABORIDE=0D Technique developed by Biology E.M. Unit Canberra=0D =0D Clean the cathode with 25% hydrochloric acid in water by immersing for 60= seconds and then cleaning with a weak alkaline (ammonia or sodium hydrox= ide). Wash with water and then alcohol before drying.=0D =0D LaB6 sources should last a long time (1000 hours plus) but they do need a= n intermediate cleaning session about every 250 to 350 hours. Some peopl= e amaze us by getting away with 1100 hours without cleaning but this is t= he exception not the rule.=0D =0D THE ELECTRON COLUMN=0D =0D The column requires cleaning when you find the stigmator controls reach t= he end of their range. In the first case change the variable aperture (f= inal aperture) to see if that enables you to carry on working within the = stigmator range. If so then you know that the aperture you first used is= too dirty for the kV you intended to use. If changing the aperture does= not change the level of astigmatism the problem is in the rest of the co= lumn; you have no alternative but to clean it.=0D =0D The column liner may be removed from underneath the anode or on some inst= ruments from the specimen chamber downwards. Some of the Hitachi range h= ave column components that are removed from the gun chamber end as well a= s the specimen area and complex manipulations may be required (watch your= service engineer).=0D =0D Clean the column liner with an Ultrasonic cleaner if possible, as a "wet"= clean is better at getting down inside the tubes. If the cleaner is amm= onia based it will attack copper-based materials so do not leave them in = the cleaner for more than a few minutes. Straight 5% ammonia solution in= water is fine but if you need a little abrasion to help the process the = commercially available "Quadralene" is ideal.=0D =0D The ultrasonic solution described in the gun cleaning process is fine pro= vided you do not leave the components in the ammonia solutions for more t= han a few minutes.=0D =0D SHINY APERTURES=0D =0D The silver coloured apertures are made of molybdenum or platinum and are = most efficiently cleaned using heat where temperatures in the orange-red = range are required.=0D =0D The ideal cleaning procedure uses a high vacuum coating unit where the ap= ertures are placed upon a platinum (for platinum) or molybdenum (for moly= bdenum) boat. Current is passed through the boat under high vacuum holdi= ng the apertures at orange-red until they display a constant colour all a= cross the aperture. Do not look at the boat without dark glasses and do = not let the temperature drift into the white range or you may melt the ap= ertures. After cleaning check with a lens that the apertures are still p= erfectly round. Throw miss shaped apertures away as they will give astig= matism problems if used.=0D =0D If you do not have a high vacuum coater you may clean platinum apertures = by holding them in a Bunsen flame using a platinum boat or platinum tippe= d tweezers. Again go to orange-red heat until the aperture glows one col= our all the way across. If you try to heat up molybdenum apertures in th= is way they will oxidise and go black. You have no alternative with thes= e apertures but to replace them.=0D =0D After cleaning any component check it thoroughly with a lens before placi= ng it back in the microscope. Always have a stock of the metal apertures= ready for replacement.=0D =0D VACUUM SEALS=0D =0D Each time you find a vacuum seal this requires some action. Remove the s= eal but ONLY USE A WOODEN STICK FOR REMOVAL. Gently pull the "O" ring th= rough the groove between the base of your thumb and first finger. This s= hould remove any debris and the "finger grease" should be sufficient to l= ubricate the "O" ring. DO NOT GREASE AN "O" RING UNLESS IT IS A MOVING S= EAL. Do not clean the "O" rings in a solvent as they will dry and start = to crack after repeated cleaning. If an "O" ring is really dirty wash it= in hot soapy water, running it between your fingers to massage in the cl= eaning media.=0D =0D Clean the "O" ring seat with a gentle solvent like alcohol before replaci= ng the seal.=0D Protrain Maintaining a Scanning Electron Microscope 3=0D =0D
} A postgraduate had prepared 3 tubes of 10g uranyl acetate in 15 mls water.
This is more UA than we usually use--our max is 1 or 2 g for 100 ml of 1% or 2% stock solutions.
} Later she decided to dispose it away. She brought it to the radiactive waste } disposal to discard it. The safety officer did a check to see whether it was } safe to dispose it there. The radiation emitted was very high, about 500 } counts per unit.
It is important to know what are "counts per unit". If you mean counts per second, that is a moderate amount of counts. It is also im- portant to know how the measurement was obtained. If a Geiger counter was used and the counts taken outside a closed jar, they likely arise from gamma rays, which will penetrate the skin and cause damage. If the counts were taken with a liquid scintillator, they could also arise from alpha particles, which are the decay product of the U, or beta particles, which are from decay of a daughter nuclide. Alphas and low-energy betas will not penetrate the dead layer of the skin. They are not harmful unless the material is inhaled, ingested or absorbed through the skin. The last process is not a problem with UA.
} She was told it was not safe for her to handle it without } protection. She was worried and went for a blood test. It showed the cell } count of lymphocyte was lower than normal. 6 weeks later she went for } another count. This time the cell count was much higher.
These counts are not specific for radiation. It is very unlikely that they are related to radiation, and much more likely that they are related to the stress from worrying about radiation. The amounts of radiation which lead to rapid changes in blood counts are massive--drin- king 30 g of UA might lead to that much radiation, and there would be chemical effects from that much UA which could cause blood cell changes.
} She is now very } worried and would like to how harmful is the radiation from uranyl acetate. } Can anybody help to ease her anxiety?
Unless the UA enters the body--as opposed to being on the skin or outside--only the gamma rays from decay of daughter products will be at all harmful. One should always wash one's hands after using UA to remove any droplets which may have gotten on the skin--especially before eating or smoking, which could lead to ingestion of the UA. The key measure for biological effects is the rem (Radiation Equivalent in Man) which is the product of the quality factor--1 for gammas & high-energy betas, about 2 for low-energy betas, and 10 to 20 for alphas--times the dose in rad (Radiation Absorbed Dose). The dose is measured in ergs per gram of tissue with 100 erg/g = 1 rad. This unit is roughly related to a mea- sure of ionization produced by a radiation field (which is measured in Roentgen units). To give an idea of possibilities for harm, the normal background radiation is about 50 mr (millirad) per year, the general population is given a limit of 500 mr per year before exposure is considered to be a problem, and radiation workers are allowed 5 rad per year. Your student should ask the safety officer how many rads were in the 30 g of UA. She should also ask how many mr/hr were measured at the outside of the jars. If this is a high number, she should calculate what she was exposed to. In particular, she should assume all the radiation would be absorbed in her hands. This calculation should reassure her.
} Till today she has not forgiven people who has been handling uranyl acetate } for not informing of the risk. } She should have been informed about the risks--everyone here who uses radioisotopes or radiation-producing equipment has to take a safety course and frequent refresher courses. UA is usually considered to be a negligable risk when used in the amounts usual for electron microscopy. There are far more serious hazards, such as OsO4 and glutaraldehyde, associated with EM. I hope this will put her mind at ease. Yours, Bill Tivol
I'm curious about using a LCTF to filter the laser. All the LCTFs I have seen have a very poor transmissivity. Doesn't this make a large impact on the spectra mwhich is collected by the CCD?
Timothy M. Prusnick 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 } -----------------------------------------------------------------------. } } The ChemIcon instrument uses the LCTF as a tunable bandpass filter. The } idea behind the instrument is to globally illuminate the sample with a large } laser spot and then collect all of the Raman scatter from the sample and } filter it through the LCTF. } } You then can collect spectra by taking a series of images with the CCD, each } at a different wavelength. If you select the same pixel in each of the } wavelength dependent images and plot it's intensity as a function of the } image wavelength, you can reproduce a spectra. Therefore, if you have a 100 } pixel by 100 pixel image, you can have 10000 spectra in one spectral image } dataset. The spectral resolution is based on the bandpass of the filter, } which is about 7 cm-1. The spatial resolution, as Fernando said, is } diffraction limited. } } Although I no longer work for the compant, I helped design the ChemIcon } instrument. So, if you have any questions about it's use or the science } behind it's design, please feel free to contact me. } } Tim Prusnick, PRUSNICK_TIM-at-worldnet.att.net } Raman Application & Support Engineer } Renishaw Inc. } 623 Cooper Court } Schaumburg, IL 60173 } PHONE: 847-843-3666 } FAX: 847-843-1744 } } -----Original Message----- } } From: Fernando Agullo-Rueda [mailto:FAR-at-icmm.csic.es] } Sent: Monday, September 14, 1998 2:35 AM } To: microscopy-at-Sparc5.Microscopy.Com } Subject: Re: Raman imaging } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } Kalpana S Katti wrote: } } } Does anayone on this network have experience with the new field of Raman } } Imaging ? We have a raman instrument with a tunable filter and imaging. I } } have a couple of questions. I am a TEM person and new at this so excuse } } me if these questions are too elementary. } } } } The manufacturer claims that on aquiring the image the spatial resolution } } of the 'spectra' is as small as a pixel. But since unlike IR, Raman is a } } scattering process wouldn't all regions limited by the beam (probe) , } } about 150 microns, be affected by each other and thus negating the claim } } of 1 pixel resolution? Infact I did observe this experimentaly while } } imaging composite polymeric samples. Spectra from all regions illuminated } } by the beam are almost identical although the image } } does exhibit contrast due to different phases. } } This question can be best answered by the manufacturer of } your instrument. However, I guess that your instrument has been } designed to get the best possible resolution. In that case, } as in other optical microscopy, the resolution is limited } by the diffraction of light. In other words, due to the wave nature } of light you can not focus a light beam below a certain size. } In the visible range that means that your spatial resolution } is around one micrometer on the sample. } } Probably the manufacturer has arranged things in such a way that } one micron on the sample is imaged at least as one pixel on the } CCD detector. } } I hope this helps. } } --------------------------------------------------------------------------- } Fernando Agullo'-Rueda } Raman Microscopy Laboratory } Instituto de Ciencia de Materiales de Madrid (CSIC) } Cantoblanco, E-28049 Madrid } Espan~a (Spain) } } Tel: +34-91-334-9015 E-mail: {FAR-at-icmm.csic.es} } Fax: +34-91-372-0623 {http://www.icmm.csic.es/} } {http://www.icmm.csic.es/Fagullo/Fagullo.htm} } ---------------------------------------------------------------------------
-- ****************************** Jim Haley Applications Engineer I-CUBE 2411 Crofton Lane, Suite 14A Crofton, MD 21114 voice: (301) 858-0505 fax: (301) 858-0615 web site: http://www.i-cubeinc.com e-mail: haley-at-i-cubeinc.com ******************************
We have an environmental SEM (ESEM model 2020) with the standard stage. It would be of great help to several of our programs to have a stage with a wider range of motions. ESEM did make and deliver stages with longer traverses. The stage we have moves only about plus or minus one inch (twenty five mm).
If you have an ESEM 2020 with the large traverse stage and are willing to part with the stage, please contact me. We would be willing to buy the stage (if a price can be agreed). We could consider exchanging our stage for yours, if the smaller motion will meet your needs.
Alwyn Eades Department of Materials Science and Engineering Lehigh University 5 East Packer Avenue Bethlehem Pennsylvannia 18015-3195 Phone 610 758 4231 Fax 610 758 4244 jae5-at-lehigh.edu
I just began using a Zeiss 10C TEM which has been used by many people before, who unfortunately are no longer here. We have a minor (?) problem with the filament, what appears to be a loose connection of some sort. When the filament is switched on the voltmeter jumps as usual to around 1.5V, but quickly drops to around 0.2-0.3V. The vacuum is fine, and so I'm led to believe it is a loose connection, or the filament has been comprimised otherwise. The service representative has not responded for a few weeks. IF anyone could offer some advice as to what exactly should be looked for in the assembly, to minimize troubleshooting time, it would be very helpful. The professor I'm working for is also new to the university and would rather to have a list of "quick options" than to take the casing apart and so forth.
TJ LaFave University of North Carolina--Charlotte Department of Physics [Department of Electrical Engineering] Charlotte, NC 28223
The Cornell Nanofabrication Facility has an open staff position for electron beam lithography and related microfabrication technologies. The position is available immediately. BS., MS, or PhD.
If you know of anyone interested, please have them contact me.
Thanks
Lynn Rathbun
***************************************************** Dr. Lynn Rathbun, User Program Manager Voice (607)-255-2329 ext 110 Cornell Nanofabrication Facility FAX(607)-255-8601 Knight Laboratory-Cornell University email Rathbun-at-cnf.cornell.edu Ithaca, New York 14853 http://www.cnf.cornell.edu/ Webmaster -at- Christian World Adoption http://www.cwa.org/ Webmaster -at- Joint Council on International Children's Services www.jcics.org (any opinions or representations of fact re: adoption are my own however) *****************************************************
Does anyone know the email address or phone number for
Vishwas Bhide ?
At one time he worked at Intel, but apparently we have lost track of him.
You can email me directly if you wish.
Thanks in advance
Fred
******************************************************** Fred Pearson Brockhouse Institute for Materials Research McMaster University 1280 Main St. West Hamilton, Ontario Canada L8S 4M1
we use chromium exclusively for coating our polymer membranes etc. for FESEM work (S900, S4500).
The thing is you MUST pump away every gas other than the sputtering gas, as Cr, unlike Au, will form nitrides & oxides which are not as dense as the metal as a coating on your specimen.
Our coater is the Xenosput, obtained through Edwards. It uses xenon as the sputtering gas. One (small) cylinder lasts ~ 10 years. The best part of its operation is that the final purging of gases in the chamber is achieved by actually sputtering titanium in the chamber itself so you only have xenon left for the final coating stage. Ours has a rotating stage and gives a very even coat over at least 75mm radius.
***************************************************** Mel Dickson, Director. Electron Microscope Unit, University of New South Wales. Sydney NSW 2052 Australia
I am new to EMs in general and have begun work on a Zeiss 10C TEM. The filament worked properly for about a week. Now, however, when the filament is activated the voltmeter immediately jumps to 1.5V (as normal), but instead of remaining there it quickly loses potential to about 0.2-0.3V. The vacuum is fine. I suspect a loose connection, but opt for an experienced person to perhaps suggest where I might begin. I am still asking around for the schematics. It shouldn't be a very difficult job (?).
Unfortunately, no one here currently knows the electronic configuration for the filament, and we prefer to know this before going through a series of unguided troubleshooting. Also, the service rep hasn't responded for some two weeks yet. Furthermore, the maintenance section of the manual is missing.
If anyone has had experience and is willing to take the time to offer some useful suggestions it would be greatly appreciated.
TJ LaFave University of North Carolina at Charlotte Department of Physics Charlotte, NC 28223 (704)547-3392
The LCTF used in the ChemIcon instrument has a transmissivity max of only 20-30%, and this can get as low as 10% throughout the free spectral range of the device. The ChemIcon instrument overcame this fault by brute force, using a much higher power laser than the more common instruments to generate the Raman scatter in experiments. In addition, the CCD itself was back thinned and had a high QE in the wavelength range of the experiment (and binning the CCD helps too - but at a loss of spatial resolution).
Tim Prusnick, PRUSNICK_TIM-at-worldnet.att.net Raman Application & Support Engineer Renishaw Inc. 623 Cooper Court Schaumburg, IL 60173 PHONE: 847-843-3666 FAX: 847-843-1744
-----Original Message----- } From: Jim Haley [mailto:haley-at-i-cubeinc.com] Sent: Monday, September 14, 1998 1:20 PM To: prusnick_tim-at-worldnet.att.net Cc: microscopy-at-Sparc5.Microscopy.Com
Timothy,
I'm curious about using a LCTF to filter the laser. All the LCTFs I have seen have a very poor transmissivity. Doesn't this make a large impact on the spectra mwhich is collected by the CCD?
Timothy M. Prusnick 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 } -----------------------------------------------------------------------. } } The ChemIcon instrument uses the LCTF as a tunable bandpass filter. The } idea behind the instrument is to globally illuminate the sample with a large } laser spot and then collect all of the Raman scatter from the sample and } filter it through the LCTF. } } You then can collect spectra by taking a series of images with the CCD, each } at a different wavelength. If you select the same pixel in each of the } wavelength dependent images and plot it's intensity as a function of the } image wavelength, you can reproduce a spectra. Therefore, if you have a 100 } pixel by 100 pixel image, you can have 10000 spectra in one spectral image } dataset. The spectral resolution is based on the bandpass of the filter, } which is about 7 cm-1. The spatial resolution, as Fernando said, is } diffraction limited. } } Although I no longer work for the compant, I helped design the ChemIcon } instrument. So, if you have any questions about it's use or the science } behind it's design, please feel free to contact me. } } Tim Prusnick, PRUSNICK_TIM-at-worldnet.att.net } Raman Application & Support Engineer } Renishaw Inc. } 623 Cooper Court } Schaumburg, IL 60173 } PHONE: 847-843-3666 } FAX: 847-843-1744 } } -----Original Message----- } } From: Fernando Agullo-Rueda [mailto:FAR-at-icmm.csic.es] } Sent: Monday, September 14, 1998 2:35 AM } To: microscopy-at-Sparc5.Microscopy.Com } Subject: Re: Raman imaging } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } Kalpana S Katti wrote: } } } Does anayone on this network have experience with the new field of Raman } } Imaging ? We have a raman instrument with a tunable filter and imaging. I } } have a couple of questions. I am a TEM person and new at this so excuse } } me if these questions are too elementary. } } } } The manufacturer claims that on aquiring the image the spatial resolution } } of the 'spectra' is as small as a pixel. But since unlike IR, Raman is a } } scattering process wouldn't all regions limited by the beam (probe) , } } about 150 microns, be affected by each other and thus negating the claim } } of 1 pixel resolution? Infact I did observe this experimentaly while } } imaging composite polymeric samples. Spectra from all regions illuminated } } by the beam are almost identical although the image } } does exhibit contrast due to different phases. } } This question can be best answered by the manufacturer of } your instrument. However, I guess that your instrument has been } designed to get the best possible resolution. In that case, } as in other optical microscopy, the resolution is limited } by the diffraction of light. In other words, due to the wave nature } of light you can not focus a light beam below a certain size. } In the visible range that means that your spatial resolution } is around one micrometer on the sample. } } Probably the manufacturer has arranged things in such a way that } one micron on the sample is imaged at least as one pixel on the } CCD detector. } } I hope this helps. } } -------------------------------------------------------------------------- - } Fernando Agullo'-Rueda } Raman Microscopy Laboratory } Instituto de Ciencia de Materiales de Madrid (CSIC) } Cantoblanco, E-28049 Madrid } Espan~a (Spain) } } Tel: +34-91-334-9015 E-mail: {FAR-at-icmm.csic.es} } Fax: +34-91-372-0623 {http://www.icmm.csic.es/} } {http://www.icmm.csic.es/Fagullo/Fagullo.htm} } -------------------------------------------------------------------------- -
-- ****************************** Jim Haley Applications Engineer I-CUBE 2411 Crofton Lane, Suite 14A Crofton, MD 21114 voice: (301) 858-0505 fax: (301) 858-0615 web site: http://www.i-cubeinc.com e-mail: haley-at-i-cubeinc.com ******************************
A solution of 10g UA in 15mls H2O was measured with a Geiger counter. } 500 counts/sec was generated. A supplier had measured 100g UA :- 1 Alpha - {2 counts/sec, using a 540 scintillation meter with AP-2 Probe 2 Beta - } 500 counts/sec, using a 540 E1 probe coupled to a GM Meter (this determines beta events and some low energy gamma events) 3 Gamma dose Rate (energy field) - two measurements done: using Mini monitor tpye R with GM Probe - 0.6mR/hr (mainly gamma) and Ionisation chamber DMM 95/0500 - 5 mR/hr (Beta and Gamma energy field). 4 Specific Activity (U approx. 55%) = 1.04 x 10 { {...} } Bq { {...} } gm { {...} } .
Can UA be used openly without protection in laboratory?
Hi Group, =20 I am looking for anybody who does or is interested in photothermal = microscopy studies. I have applied photothermal techniques for optical microscopy = to enhance optical sensitivity during investigation of living cells. = However I have found very little info about the subject and I would like to exchange = with ideas with somebody who has similiar experience/interest. =20 Dmitri Lapotko =20 Luikov Heat and Mass Transfer Institute 15 Brovka Street Minsk Belarus =20 tel:(375172)842483 fax:(375172)842486 e-mail: ld-at-ns1.hmti.ac.by
} With my tongue only partway in my cheek, I ask... Alconox is a *mild* } detergent? What do you consider to be a harsh one? } I know no detergent is going to attack stainless, but read the fine } print on the packages before you try soaking on aluminum & its alloys, } some are alkaline enough to etch the surface. } Dear Robert, I'm doing the experiment. I mixed up a 1% solution of Alconox per the directions on the package. The pH is 9.1, so this is not too alkaline a detergent. At present, there is a piece of Al foil in the solution; I'll let you & the list know what fate the foil suffers. Yours, Bill Tivol
Hi everyone! My computer has been upgraded. My E-mail address has been changed to reflect my married name. New address is : brygv-at-ferro.com thanks, Vicky
} ------------------------------------------------------------------------ } 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, } Has anyone experienced puckering of tissues embedded in LRWhite resin? My tissues are cell cultures fixed in 2%PFA, .5%GA, dehydrated to75% ETOH then into graded LRWhite resin. I cut on a diamond knife and embed on 200 mesh grids. Before staining, the puckers and wrinkles are seen as small, and frequen, folds over most cells or along the cell borders. The resin areas are totally free of folding. It's as if the tissue area is picking up H2O during sectioning then has no where to go when it re-dries....very frustrating. After staining it's worse, as the stain seems to stay in the folds and gets really dark. Any thoughts and suggestions are very welcome as always. } Linda Fox } Loyola University } Stritch School of Medicine } lfox1-at-wpo.it.luc.edu } } Hi, You have several problems here. First, acrylics like LR White, etc., do not bond WITH the tissue as do epoxies. This allows a lot of shifting as soon as the stresses are relieved by cutting a thin section.. LR White also is modestly poorly crosslinked (as compared to the epoxides). You can increase the crosslinkage chemically (unless you are doing immunostaining, then you do not want to heavily crosslink). What to do? Do you need to use LR White? What is your purpose of using it. It has a number of downsides which one can trade off in the immunoprocessing protocols for better location of antigens. For standard TEM work is is inferior to epoxides (wrinkling of thick sections, poor crosslinkage, polymerization damage, difficulty sectioning because of its property of attracting water to the block face, etc.) What to do? If you must use LR White, try using a 100 mesh grid with a film. Also when picking up floating grids, DO NOT suck the water off under the grid with filter paper. Do draw the water off by sticking filter paper between the forceps. Then, lay the grid in the forcepts down to dry. Pull a lamp with a 60W bulb down over the forceps to slightly warm the situation. Use at least 5 forceps at once. That way some grids are drying and others are being picked up. This is the recommendation of Newman who holds the patent for LR White and LR Gold (He sold the license to the London Resin Company.) Should you decide to more heavily crosslink the resin, contact EMS for advice and the chemical. (I have no stock in EMS). If your sections have folds and you posstain them with heavy metals, the stain will accumulate in the folds. There is nothing you can do about that. You must avoid the wrinkles in the first place. Should you have more trouble, contact me. Bye, Hildy
An addendum to Melvyn Dickson's message, particularly for our younger readers:
When I started here at Reading, there was still in operation an EM6 microscope (AEI) which used a lot of what we Britishers call "valves" and Americans call "tubes" (what about Australians?). These all had an iridescent shine from the little bit of metal (magnesium?) which has been fired to scavenge the last traces of gas, a process known as GETTERING.
} Our coater is the Xenosput, obtained through Edwards. It uses xenon as the } sputtering gas. One (small) cylinder lasts ~ 10 years. The best part of } its operation is that the final purging of gases in the chamber is achieved } by actually sputtering titanium in the chamber itself so you only have } xenon left for the final coating stage.
Concerning our Korean friend's enquiry about phase contrast, I was at a Scanning Force Microscopy workshop in Bristol last week. Prof. Andrew Keller gave a most interesting historical talk, suggesting that the new technique would follow a three stage development similar to that of Electron Microscopy:
(1) ELATION: The EM allowed one to visualize a dimension previously only inferred from colloid studies;
(2) ANTICLIMAX: it looked at the surface only, and was prone to artefacts;
(3) STEADY PROVEN USEFULNESS.
Hence the importance of backing up with other techniques, particularly optical microscopy. As regards polymers, what pulled EM out of the doldrums in the late '50s was being able to see the same polymer crystals (solution grown polyethylene lamellae) both under the TEM and the newly available Zernicke Phase Contrast microscopes.
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
ELECTRON MICROSCOPY RESEARCH TECHNICIAN POSITION AVAILABLE
Laboratory of William Lehman Ph. D. Department of Physiology Boston University School of Medicine 80 East Concord Street Boston, MA 02118
Qualifications:
1. Experience in molecular electron microscopy, particularly cryo-EM. 2. Facility with computer based-image analysis. 3. Familiarity with handling purified protein samples. 4. Ability to work independently in a small group environment.
Brief description of project:
We carry out state of the art - high resolution electron microscopy, computer assisted image analysis and three-dimensional image reconstruction to determine the arrangement of muscle thin filament components on F-actin and evaluate their position and influence on actin domains. Many significant projects, both on smooth and skeletal muscle systems, are being performed, and there is an excellent opportunity for a motivated individual to contribute to our understanding of muscle contractility and regulation.
Some recent publications:
Lehman, W., R. Craig & P. Vibert (1994) Ca-induced tropomyosin movement in Limulus thin filaments revealed by three-dimensional reconstruction. Nature 368, 65-67.
Lehman, W., P. Vibert, P. Uman & R. Craig (1995) Steric-blocking by tropomyosin visualized in relaxed vertebrate muscle thin filaments. J. Mol. Biol. 252, 191-196.
Vibert, P., R. Craig & W. Lehman (1997) Steric-model for activation of muscle thin filaments. J. Mol. Biol. 266, 8-14.
Hodgkinson, J.L., M. EL-Mezgueldi, R. Craig, P. Vibert, S.B. Marston & W. Lehman (1997) 3-D Image reconstruction of reconstituted smooth muscle thin filaments containing calponin: Visualization of interactions between F-actin and calponin. J. Mol. Biol. 273, 150-159.
Lehman, W., P. Vibert & R. Craig (1997) Visualization of caldesmon on smooth muscle thin filaments. J. Mol. Biol. 274, 310-317.
Hanein, D., N. Volkmann, Goldsmith, S., A.-M. Michon, W. Lehman, R. Craig, D. DeRosier, S. Almo & P. Matsudaira (1998) An atomic model of fimbrin binding to F-actin and its implications for filament crosslinking and regulation. Nature Struct. Biol. 5 787-792.
Send resume and 3 references to Dr. Lehman at above address or either FAX to (617)638-4273 or e-mail to lehman-at-med-rana.bu.edu.
We ion beam sputter Pt for polymer FESEM work up to ~X100,000; Pt gives us no significant coating artifacts up to that magnification and, unlike Cr, doesn't require pre-sputtering the target to remove the native oxide. For higher mags we will Cr coat.
Ev Osten 3M Company St. Paul, MN efosten-at-mmm.com
Stephen McCartney {stmccart-at-vt.edu} on 09/14/98 10:29:47 AM
To: Microscopy-at-sparc5.microscopy.com cc: (bcc: Ev Osten/US-Corporate/3M/US)
I am interested in high resolution coaters to use with field emission SEM. Does anyone working with FE-SEM's have some opinions for me on what is best specifically for polymer applications. Steve
------------------------------ Stephen McCartney Research Associate Virginia Tech Materials Institute 2108 Hahn Hall Blacksburg, VA 24061-0344 USA
HELP! I am new to the microscopy listserver and I am currently conducting undergraduate research for the Department of Biomedical Sciences at Southwest Missouri State University, located in Springfield, Missouri (USA). I am attempting to develop a TEM protocol to study the gap junction protein, Connexin-43 (Cx43) at the ultrastructural level. I am specifically interested in a protocol that utilizes colloidal gold as a marker and a post embedding technique that uses conventional resins. If anyone would be willing to share a specimen protocol for the localization of Cx43 or any other Connexins, please reply to me off list at: aaronrea-at-hotmail.com Thank you very much for your time!
Aaron Rea Department of Biomedical Sciences Southwest Missouri State University aaronrea-at-hotmail.com
I have some experience in evaporating silver, (2 - 3mm shot) from a tungsten wire basket, however, I am now faced with challenge of evaporating Au wire onto a similar substrate. Is it appropriate to simply wrap the Au wire around the larger diameter tungsten wire and proceed or is there a better approach. Your comments/suggestions are appreciated. Thanks.
Regards, Paul Gerroir Xerox Research Center of Canada
The Microscopy and Microanalysis Center (MMC) at the University of = Maryland at College Park is searching for a laboratory manager to = maintain and run two TEMs (Hitachi 600 AB, and JEOL 4000FX), one = electron microprobe (JEOL 8900), an environmental SEM (Electroscan E3) = and AFM (Dimension 3000) among other equipment. The MMC is a campus = facility that provides service to faculty, students, and outside users. = The facility is also used for teaching and research. The laboratory = manager would supervise two graduate students in the use of the = equipment.
Qualifications: The qualified candidate should have experience in the = maintenance of electron microscopes and their use. Background on = electronics and vacuum technology is required. Bachelor or Masters = degree in Materials Science, Physics or Engineering is preferred. Good = oral and written skills, as well as supervisory skills are required. =20
Salary: The starting salary is $30,000 to $35,000 depending on = experience. =20
Availability: October 1, 1998.
For best consideration, interested candidates should send curriculum = vitae and a list of three references to:=20 Dr. Lourdes Salamanca-Riba at either riba-at-eng.umd.edu,=20 Fax No. (301) 314-9467, or=20 Materials and Nuclear Engineering Department University of Maryland College Park, MD 20742-2115
The University of Maryland is an equal opportunity affirmative action = employer.
Are any of you interested in the possibility of using your micrographs to illustrate children's books? Here's a conference that should be quite informative:
Marine Biological Laboratory Hosts Institute for Children's Book Authors and Illustrators October 9-11, 1998
Woods Hole, MA-The Marine Biological Laboratory's Science Writing Fellowships Program and the Center for Children's Environmental Literature is co-sponsoring an Author, Illustrator, Biologist Institute. Working and aspiring children's book authors and illustrators, as well as scientists, are invited to participate in the three-day meeting. Organizers hope to foster new collaborations between authors, illustrators, and biologists.
For more information, contact: Pamela Clapp Hinkle Director of Communications Marine Biological Laboratory 7 MBL Street Woods Hole, MA 02543 Tel: 508-289-7276 Fax: 508-457-1924 e-mail: pclapp-at-mbl.edu
For a complete program, see http://www.mbl.edu/html/MISC/AIB.html.
Caroline Schooley Educational Outreach 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/Books.html Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/pci.html
In a message dated 98-09-15 19:07:25 EDT, you write: { { I have some experience in evaporating silver, (2 - 3mm shot) from a tungsten wire basket, however, I am now faced with challenge of evaporating Au wire onto a similar substrate. Is it appropriate to simply wrap the Au wire around the larger diameter tungsten wire and proceed or is there a better approach. Your comments/suggestions are appreciated. Thanks.
Regards, Paul Gerroir Xerox Research Center of Canada } }
Yes - it works well to wrap the wire around the tungsten filament.
A word of caution on the evaporation: Heat up the filament slowly, observing the gold through suitable dark glasses. At some point the gold will melt and run into a ball in the "v" of the filament. Then you can turn up the filament current until the gold evaporates.
Hello! I work with Atriplex nummularia Lidl. and this specie has a profusion of vesicular trichomes on both surfaces that are very fragile. I'm trying to obtain a transversal view of the leaf lamina to measure the thickness of the trichome layer.=20 Who has any idea? Thanks in advance. Rejane
Rejane Magalh=E3es Pimentel Galindo =20 ggalindo-at-elogica.com.br Universidade Federal Rural de Pernambuco Av. Boa Viagem, 6592/602 FAX: 55 (081) 4416177 51130-000, Recife, Pernambuco, Brasil
At 03:57 PM 9/15/98 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Paul,
I once watched someone hang a piece of wire from a larger tungsten wire, letting it dangle by a loop (like a clothes hangar). While every else predicted that the metal would simply fall off the wire when the loop heated up, the technician simply increased the current until the wire melted, forming a perfect adhering drop. He then increased the current until the metal evaporated and got a beautiful coating on his substrate. I don't remember if it was Au, Ag, or Pt, but the technique was simple, and it worked for him. Until then, we had been carefully wrapping the wire to evaporate around the supporting wire, like an electrical coil---a real skill for some of us.
For what it's worth.
Randy
} } Randy Tindall 2017 Princess Jeanne Las Cruces, New Mexico 88001-4157
The concentration quoted is far higher (btw at what concentration in water does UA become a saturated solution?) than normally would be used for EM staining.
} A solution of 10g UA in 15mls H2O was measured with a Geiger counter.
I have been using UA for many years and I recall that whenever I questioned and investigated its possible radiation implications I have been assured that it is not dangerous at the concentrations and quantities we use, provided that it is not ingested. Robin H Cross Director : EM Unit, Rhodes University, Grahamstown, South Africa eurc-at-giraffe.ru.ac.za - tel: +27 46 603 8168 - fax: +27 46 622 4377 http://www.ru.ac.za/affiliates/emu/em.htm
I can second what Randy says. I have been evaporating gold wire off larger diameter tungsten wire (occasionally) for 30 years. It works, in an old Edwards coating unit. I must admit, I have always wrapped the gold around the tungsten, using two pairs of clean forceps.
Ann Fook Yang EM Unit Eastern Cereal and Oilseed Research Centre Agriculture and Agri-Food Canada 960 Carling Ave Central Experimental Farm Ottawa, Ontario Canada K1A 0C6
The concentration quoted is far higher (btw at what concentration in water does UA become a saturated solution?) than normally would be used for EM staining.
} A solution of 10g UA in 15mls H2O was measured with a Geiger counter.
I have been using UA for many years and I recall that whenever I questioned and investigated its possible radiation implications I have been assured that it is not dangerous at the concentrations and quantities we use, provided that it is not ingested. Robin H Cross Director : EM Unit, Rhodes University, Grahamstown, South Africa eurc-at-giraffe.ru.ac.za - tel: +27 46 603 8168 - fax: +27 46 622 4377 http://www.ru.ac.za/affiliates/emu/em.htm
I used to evaporate Au in a tungsten basket in an old Denton evap coater. We simply took a length of Au wire, wadded it up into a tiny ball and placed that in the basket. Took no large amount of manual dexterity and gave good results.
Gerroir, Paul J wrote:
} Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
Dear Josephine, } } A solution of 10g UA in 15mls H2O was measured with a Geiger counter. } 500 } counts/sec was generated. } A supplier had measured 100g UA :- } 1 Alpha - {2 counts/sec, using a 540 scintillation meter with AP-2 Probe } 2 Beta - } 500 counts/sec, using a 540 E1 probe coupled to a GM Meter (this } determines beta events and some low energy gamma events) } 3 Gamma dose Rate (energy field) - two measurements done: } using Mini monitor tpye R with GM Probe - 0.6mR/hr (mainly gamma) } and Ionisation chamber DMM 95/0500 - 5 mR/hr (Beta and Gamma energy } field). } 4 Specific Activity (U approx. 55%) = 1.04 x 10 { {...} } Bq { {...} } gm } { {...} } . } I calculated approximately the expected activity from 30 g UA (about 0.1 mole, or 6*10^22 atoms). T_1/2 is 4.4*10^9 y and there are 3.1*10^7 s/y, so the decay rate is 5*10^-18 s^-1, and the activity is 3*10^5 Bq. The build-up of daughter products with shorter half-lives will reach steady state at which point the activities of the daughters will be the same as that of the parent. Pa 234 has a gamma transition, and there are several betas in the chain. The longer-lived isotopes in the chain have lives of 10^4 to 10^5 y, and these will not be at steady state (unless your UA is *very* old ;-) ). 0.6 mR/hr is a significant amount of exposure, and, if one were to hold the jars for some minutes, a sizable fraction of the allowed annual dose would be attained.
} Can UA be used openly without protection in laboratory? } Small amounts can be used, but be sure to wash hands before eating. One area of the lab should be used for UA. A quiet area with little traffic is best. UA, while not nearly the most dangerous EM reagent, should still be treated with respect. Yours, Bill Tivol
Responding to the message of {01J1VEY9MEFC8WXL78-at-pt.Cyanamid.COM} from "corwinl-at-pt.cyanamid.com"-at-Sparc5.Microscopy.Com: }
} According to the Handbook of Chemistry and Physics, the solubility of } UO2(C2H3O2)2.2H2O is 7.694 g/100 mL in 15 deg C water. } } } Leonard Corwin } Fort Dodge Animal Health } Princeton, NJ 08543-0400
That's just the information I was looking for!
In thinking about my own experiences with solubility of UA, I've noticed that even at only 2, 3, 4 % w/v, there always seems to be a bit that doesn't go into solution, even with heating. I consulted my vendor of UA and was told that there is an insoluble component present in commercial UA, and thats what I was seeing. This insoluble contaminant would then confuse efforts to determine saturation point by visual obvservation - are those insoluble contaminants or are they leftover UA crystals, having reached saturation? The yellow color of the solution foils any attempt to detect differences in color of the two kinds of crystals.
I did isolate these unknown crystals (forgot what color they were), did EDS analysis on 10 clumps of crystals and found that in addition to moderate to high amounts of U (quick, on-the-fly subjective semi-quant intuition), there were also high amounts of titanium (one very high), moderate to high amounts of silicon (one very high), low amount of aluminum (not always present), low amount of iron (always present), varying carbon and oxygen, low phosphorus. One cubic crystal was quite high in Si and Ti, only.
As U has one of the highest backscatter coefficients, perhaps the Ti, Fe, Al is coming from stray x-rays generated inside the SEM chamber by BSE's. If not, then my results indicate the basic composition of the contaminant crystals. Any idea what they might be?
I should now take UA crystals out of the bottle, measure their spectrum, to compare with the "contaminants" spectra measured above to sort this out.
But thanks to the above solubility data, we can at least make saturated solution of UA, and ignore or filter out the contaminant crystals.
Gib Ahlstrand Electron Optical Facility, University of Minnesota, Dept. Plant Pathology 495 Borlaug Hall, St. Paul, MN. USA. 55108 (612)625-8249 612-625-9728 FAX, giba-at-puccini.crl.umn.edu
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
} EDX folks: } } Does anyone know of or plan to pick up support and/or development for } NIST's Desktop Spectrum Analyzer (DTSA) software?
At a DTSA workshop a couple of years past, Bob Myklebust *hinted* that he might start a private consulting business after retiring from NIST that would, among other things, provide DTSA training and support. But I haven't heard any announcements.
Now that DTSA has been "open sourced" by NIST, perhaps we need to start up a mailing list for user support. ( I'm CC-ing this to the microprobe list, which might be a more appropriate place for DTSA discussions until such a list can be set up. )
As long as you brought up this topic: Can anyone explain what exactly were the circumstances behind NIST's decision ? I got one semi-official explaination, which didn't exactly jive with either the currently posted comments on NIST's web pages nor with various rumours about law-suits that I heard from various vendors. Anyone know the story?
---| Steven D. Majewski (804-982-0831) {sdm7g-at-Virginia.EDU} |--- ---| Department of Molecular Physiology and Biological Physics |--- ---| University of Virginia Health Sciences Center |--- ---| P.O. Box 10011 Charlottesville, VA 22906-0011 |---
"I'm not as big a fool as I used to be, I'm a smaller fool." - Jack Kerouac Some of the Dharma {http://members.aol.com/kerouacsis/SomeDharma.html}
If you have Zephyr (model ZEM 300SW) water chillers and have experienced cooling/heating problems, please let me know how the problems were solved. One Zephyr won't cool-- water temp is 25 C. The other won't heat--temp is 8 C. Our A/C people have always been able to solve Haskris and Neslab chiller problems but are unable to solve Zephyr problems. Suggestions?
John C. Wheatley Lab Manager Arizona State University Center for Solid State Science PSA-213 BOX 871704 Tempe, AZ 85287-1704
The Midwest Microscopy and Microanalysis Society will host a Materials Sciences meeting on October 9, 1998, at Purdue University. The meeting= has been organized by Eric Kvam and includes the following sessions:
9:00 Welcome and Introduction 9:20 Advances in Scanning Force Microscopy, R.P. Andres 10:00 High Resolution EELS on Superconductor Grain Boundaries, J.L. Lee=
10:20 Coffee Break 10:30 ESEM In situ Observation of Fatigue Crack Propagation, B.M. Hillb= erry 11:10 Electron Beam Lithography using the SEM, D.B. Janes 11:30 Lunch 1:00 Laboratories Tour 2:30 People to People: Electron Microscopy in China, J.L. Lee 2:50 Coffee Break 3:10 TEM of Electronic Composite Films, D.E. Collins 3:30 Advanced Image Analysis of Concrete Microstructures, S. Diamond 3:50 Dislocation Structure and Electronics in Semiconductors, V. Gopal
Pre-registration is NOT required, and there is no registration fee for members of MMMS. For non-members who wish to attend, it will be possi= ble to join the society at the meeting. Dues are $10 for regular members and = $5 for student members. Current MMMS members will receive a complete agenda a= nd travel information in the mail next week. Others may contact me via e-= mail at jane.a.fagerland-at-abbott.com, and meeting information will be mailed = or faxed to you.
Jane A. Fagerland, Ph.D MMMS Dept. Microscopy and Microanalysis Abbott Laboratories Abbott Park IL 60064 (847) 935-0104 =
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Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
I would like to fix small spiders for TEM. I am not particular interested in preservation of specific structures, but would like to preserve both integumental and inner structures in general. I noticed that my specimens floated, and I am afraid that the fixation will not penetrate properly.
Does anybody has experience with fixation of similar specimens ? I would appreciate your comments very much.
Thank you.
Regards
Peter Funch
______________________________________________________________________ Peter Funch Assistant Professor, Ph.D.
Department of Zoology Direct Line + 45 8942 2764 Institute of Biological Sciences Secretary + 45 8942 2727 University of Aarhus Telefax + 45 86 12 51 75 Universitetsparken E-mail: peter.funch-at-biology.aau.dk Building 135 DK-8000 Aarhus C Denmark ______________________________________________________________________
Department of Cell and Structural Biology University of Illinois at Urbana Champaign ---------------------------------------------------------------------------
Qualifications: Minimum requirement: Bachelor of Science degree. Applicants with advance degrees are invited to apply. Previous lab and/or electron microscopy experience required.
Responsibilities: To perform research in a laboratory of cell biology and structural biology. Work will combine electron microscopy and light microscopy with tissue culture, immunostaining, and molecular biology.
Salary: Dependent on qualifications.
Starting Date: As soon as possible after closing date.
Send Applications to: Ms. Helen Neef (asb) Department of Cell and Structural Biology B107 Chemical and Life Sciences Laboratory University of Illinois 601 S. Goodwin Avenue Urbana, IL 61801 Phone: (217) 244-6638
Closing Date: Nov. 15, 1998. Interviews may be conducted before the closing date, but all applications received by that date will receive full consideration and the final selection will not be made until after that date.
UNIVERSITY OF ILLINOIS AT URBANA-CHAMPAIGN IS AN AFFIRMATIVE ACTION, EQUAL OPPORTUNITY EMPLOYER
****************************************************** Andrew Belmont 217-244-1648 (fax) Associate Professor 217-244-2311 (office) Department of Cell and Structural Biology asbel-at-uiuc.edu University of Illinois, Urbana-Champaign B107, 601 S. Goodwin Ave. Urbana, IL 61801 ******************************************************
Hi, I have a request for a micrograph of the pits and lands on a music CD. I thought this would be fun and simple but we aren't having any luck imaging anything (actually it was so reflective we imaged the detector!:-) Here's what we've done: We cut up a CD - using a piece of it close to the center hole (so we knew it had something on it to see). The piece was mounted (lower surface up) on a stub using a carbon sticky tab and silver paste. The sample was sputter coated for 2 minutes (some for 3) and scoped. The first time we viewed it we didn't coat it (see paragraph one for details).
Do we need a solvent to munch awhile on the surface layer of the CD? Any help would be greatly appreciated!
It was a Christmas music CD so maybe that is the problem :-).
beth
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
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Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
Numbers of JEOL owners in Oz have complained of similar oil contamination. When we last evaluated FESEMs the JEOL rep was persistent to know why we didn't like the JEOL models. One of my reasons was the oil contamination problem and I asked for some explanation as to why it was so common.
His answer was that users were opening the inner door of the airlock too soon and perhaps too quickly. There was too much air left in the airlock and the pressure rise in the chamber stalled the diffusion pump and backstreaming of the diffusion pump oil caused the contamination.
***************************************************** Mel Dickson, Director. Electron Microscope Unit, University of New South Wales. Sydney NSW 2052 Australia
Hi all, I was just wondering why everyone uses gold wire? Last time I looked, 5 9's gold splatters were the cheapest. With wire and shot you have to pay for the manufacturing/molding costs. I have always used a tungstun or moly boat, so there is no need to wrap or hang anything. -regards -andrew
--ListServer-at-MSA.Microscopy.Com
On Wed, 16 Sep 1998 10:02:47 Keith Ryan wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-----== Sent via Deja News, The Discussion Network ==----- http://www.dejanews.com/ Easy access to 50,000+ discussion forums
Hi all, I was just wondering why everyone uses gold wire? Last time I looked, 5 9's gold splatters were the cheapest. With wire and shot you have to pay for the manufacturing/molding costs. I have always used a tungstun or moly boat, so there is no need to wrap or hang anything. -regards -andrew
--ListServer-at-MSA.Microscopy.Com
On Wed, 16 Sep 1998 10:02:47 Keith Ryan wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-----== Sent via Deja News, The Discussion Network ==----- http://www.dejanews.com/ Easy access to 50,000+ discussion forums
Hi all, I was just wondering why everyone uses gold wire? Last time I looked, 5 9's gold splatters were the cheapest. With wire and shot you have to pay for the manufacturing/molding costs. I have always used a tungstun or moly boat, so there is no need to wrap or hang anything. -regards -andrew
--ListServer-at-MSA.Microscopy.Com
On Wed, 16 Sep 1998 10:02:47 Keith Ryan wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
-----== Sent via Deja News, The Discussion Network ==----- http://www.dejanews.com/ Easy access to 50,000+ discussion forums
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
Hi all, I was just wondering why everyone uses gold wire? Last time I looked, 5 9's gold splatters were the cheapest. With wire and shot you have to pay for the manufacturing/molding costs. I have always used a tungstun or moly boat, so there is no need to wrap or hang anything. -regards -andrew
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On Wed, 16 Sep 1998 10:02:47 Keith Ryan wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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Hi all, I was just wondering why everyone uses gold wire? Last time I looked, 5 9's gold splatters were the cheapest. With wire and shot you have to pay for the manufacturing/molding costs. I have always used a tungstun or moly boat, so there is no need to wrap or hang anything. -regards -andrew
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On Wed, 16 Sep 1998 10:02:47 Keith Ryan wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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Hi Group, I am looking for anybody who does or is interested in photothermal microscopy studies. I have applied photothermal techniques for optical microscopy to enhance optical sensitivity during investigation of living cells. However I have found very little info about the subject and I would like to exchange with ideas with somebody who has similiar experience/interest. Dmitri Lapotko Luikov Heat and Mass Transfer Institute 15 Brovka Street Minsk Belarus tel:(375172)842483 fax:(375172)842486 e-mail: ld-at-ns1.hmti.ac.by
In a message dated 98-09-16 20:01:57 EDT, you write:
{ { Hi, I have a request for a micrograph of the pits and lands on a music CD. I thought this would be fun and simple but we aren't having any luck imaging anything (actually it was so reflective we imaged the detector!:-) Here's what we've done: We cut up a CD - using a piece of it close to the center hole (so we knew it had something on it to see). The piece was mounted (lower surface up) on a stub using a carbon sticky tab and silver paste. The sample was sputter coated for 2 minutes (some for 3) and scoped. The first time we viewed it we didn't coat it (see paragraph one for details).
Do we need a solvent to munch awhile on the surface layer of the CD? Any help would be greatly appreciated!
It was a Christmas music CD so maybe that is the problem :-).
beth } }
How about making a replica and imaging it in the TEM?
If you wish to do that I can give you some technique hints.
I have been asked to compare the grain size of 2 MgO aggregates. The samples were embedded and polished with diamond paste, but I am not happy with the result. The surface looks smeared. Does the sample need to be etched or do I need to evaluate my polishing technique???? Thanks for you help.
Nan Laudenslager Specialty Minerals, Inc. Easton, PA nhl-at-early.com
=46or Lorentz microscopy purposes we have modified a 200CX such that specime= n is located in a non-traditional position of the column. For the specimen to be in focus the objective lens is operating at a level that is 20% higher than normal. We would like to determine the Cs of the objective lens under these conditions. Note: the image resolution is about 50=C5 under these conditions. Any ideas??
Thanks,
Mark A. Wall L-350 7000 east Ave Chemistry & Materials Science Dept. Lawrence Livermore National Lab Livermore, CA 94550 925 423-7162
All the work we did in the past proved to us beyond doubt that the oil wa= s from the rotary pump. No matter how sophisticated the system is if you u= se a rotary pump at all in the cycle you do seem to get RP oil as a contaminant.
I think the guy had a good try but I for one do not believe him!
Hi all. Great topic. We have been involved in building a new control unit for a factory which = evaporates aluminium onto plastic parts to give them that silver look. This has given us a lot more understanding of the coating technique as = they have very large vacuum chambers and many samples that need an even = and smooth coat each time. What they do is to use tungsten wires coils that have two strands of W = wire twisted together as aposed to, what we all seem to use, the single = strand of W wire. The aluminium wire is then twisted by hand onto this = coil, fairly loosely but just that it makes contact. The reasoning is = that as you start the heating of the aluminium it will melt and flow, = via capillary action, between the two W wires. This means that the = aluminium is then in very good contact with the W coil and a lower = current is needed to vaporise the aluminium. You also have a source of = aluminium the whole length of the W coil each time. This ensures a even = and repeatable coat each time.
Cheers Luc Harmsen=09 Anaspec, South Africa International technical support on microscopy. Tel: +27 (0) 11 476 3455 Fax:+27 (0) 11 476 7290 anaspec-at-icon.co.za
-----Original Message----- } From: Timothy Moeller [SMTP:tmoeller-at-noran.com] Sent: Wednesday, September 16, 1998 7:15 PM To: Paul.Gerroir-at-crt.xerox.com Cc: Microscopy-at-sparc5.microscopy.com; Microscopy-at-sparc5.microscopy.com
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Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } =20 } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of = evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the = Au } wire around the larger diameter tungsten wire and proceed or is there = a } better approach. Your comments/suggestions are appreciated. Thanks. } =20 } Regards, } Paul Gerroir } Xerox Research Center of Canada
--=20 ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
Jon, In the message below, I found a reference to a microprobe listserver. You should check it out. The website is at http://www.microanalysis.org/mas/maslistserver/maslistserver.html
This mailing list is sponsored by the Microbeam Analysis Society and managed by John Mansfield and Greg Meeker. More information is available at at the MAS web. To subscribe send email to the mailing list at:
microprobe-at-www.microanalysis.org with the word "subscribe" in the subject.
Hope this is as useful as the microscopy listserver is to me.
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On Wed, 16 Sep 1998, Heeschen, Bill (WA) wrote:
} EDX folks: } } Does anyone know of or plan to pick up support and/or development for } NIST's Desktop Spectrum Analyzer (DTSA) software?
At a DTSA workshop a couple of years past, Bob Myklebust *hinted* that he might start a private consulting business after retiring from NIST that would, among other things, provide DTSA training and support. But I haven't heard any announcements.
Now that DTSA has been "open sourced" by NIST, perhaps we need to start up a mailing list for user support. ( I'm CC-ing this to the microprobe list, which might be a more appropriate place for DTSA discussions until such a list can be set up. )
As long as you brought up this topic: Can anyone explain what exactly were the circumstances behind NIST's decision ? I got one semi-official explaination, which didn't exactly jive with either the currently posted comments on NIST's web pages nor with various rumours about law-suits that I heard from various vendors. Anyone know the story?
---| Steven D. Majewski (804-982-0831) {sdm7g-at-Virginia.EDU} |--- ---| Department of Molecular Physiology and Biological Physics |--- ---| University of Virginia Health Sciences Center |--- ---| P.O. Box 10011 Charlottesville, VA 22906-0011 |---
"I'm not as big a fool as I used to be, I'm a smaller fool." - Jack Kerouac Some of the Dharma {http://members.aol.com/kerouacsis/SomeDharma.html}
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Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
On 16 Sep 98 at 16:52, The slowly moving finger of Beth Richardson 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 have a request for a micrograph of the pits and lands on a music } CD. I thought this would be fun and simple but we aren't having any } luck imaging anything (actually it was so reflective we imaged the } detector!:-) Here's what we've done: We cut up a CD - using a piece } of it close to the center hole (so we knew it had something on it to } see). The piece was mounted (lower surface up) on a stub using a } carbon sticky tab and silver paste. The sample was sputter coated } for 2 minutes (some for 3) and scoped. The first time we viewed it } we didn't coat it (see paragraph one for details). } } Do we need a solvent to munch awhile on the surface layer of the CD? } Any help would be greatly appreciated! } } It was a Christmas music CD so maybe that is the problem :-). } } beth } Beth I have done this in the past using a rewritable CD where the reflective coating can be readily stripped using tape. Using a normal CD I think we had to physically rip it apart, ( I seem to remember it was quite tough ), a good source for this is the freebies that you find on computer mags. John John Findlay Science Faculty EM Facility. Edinburgh University. Daniel Rutherford Bldg. Kings Buildings. Edinburgh EH9 3JH. tel. 0131-650-5344 fax. 0131-650-6563 John.Findlay-at-ed.ac.uk
this may be of no help what-so-ever, but we have a Flowcool water cooler. It has a digital readout of temperature and at present it 'thinks' that the water is about 70 deg C (ie temp readout is a bout 70) whereas the true temperature on the microscope is between 16 to 18 deg C.
This has happened once before and it turned out that it was the temperature sensor which was easy and fairly cheap to replace - so we did it and it worked. It looks like it's gone again (not very reliable) after 3 or 4 years and will be replaced soon.
You might just have a similar problem.
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk ---------- } From: John C. Wheatley To: Microscopy
If you have Zephyr (model ZEM 300SW) water chillers and have experienced cooling/heating problems, please let me know how the problems were solved. One Zephyr won't cool-- water temp is 25 C. The other won't heat--temp is 8 C. Our A/C people have always been able to solve Haskris and Neslab chiller problems but are unable to solve Zephyr problems. Suggestions?
John C. Wheatley Lab Manager Arizona State University Center for Solid State Science PSA-213 BOX 871704 Tempe, AZ 85287-1704
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Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
The pits and lands on a CD are { {inside} } the plastic of the CD, so you need to dissolve it with some sort of solvent (I believe I used methanol, but can't recall for sure). The correct organic solvent will make the plastic disappear entirely, not just craze it and turn it cloudy. I think there may be several types of plastic used as the base, because I used toluene a number of years ago the first time I tried this, and it didn't work on the CD I used a few months ago. Anyway, use a couple of fresh changes until the foil is floating free, then mount it on a stub and take a look. You might need to mount both sides of a single piece so that you are sure you have the side with the pits. If you've gotten all the plastic off, you might get away without coating, but I coated mine anyway.
You can see an example of the pits and lands at http://www.mta.ca/~jehrman/cd.htm BTW, the CD I used is a Microsoft(TM) Office demo CD, so any flaws are Bill Gate's fault, not mine!
Cheers,
Jim
--
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
} ------------------------------------------------------------------------ } } 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 request for a micrograph of the pits and lands on a music CD. } I } thought this would be fun and simple but we aren't having any luck } imaging } anything (actually it was so reflective we imaged the detector!:-) } Here's what we've done: } We cut up a CD - using a piece of it close to the center hole (so we } knew } it had something on it to see). The piece was mounted (lower surface } up) on } a stub using a carbon sticky tab and silver paste. The sample was } sputter } coated for 2 minutes (some for 3) and scoped. The first time we viewed } it } we didn't coat it (see paragraph one for details). } } Do we need a solvent to munch awhile on the surface layer of the CD? } Any help would be greatly appreciated! } } It was a Christmas music CD so maybe that is the problem :-). } } beth } } ************************************** } Beth Richardson } EM Lab Coordinator } Botany Department } University of Georgia } Athens, GA 30602 } } Phone - (706) 542-1790 } FAX - (706) 542-1805 } Email - beth-at-dogwood.botany.uga.edu } **************************************
There was a thread about CDs recently which might be helpful. In a mass produced CD the pits are in the aluminum layer which is under a plastic layer. Electron imaging won't stand a chance unles the plastic is removed. BTW... CD-Rs are quite different - use a dye rather than Al film.
Woody McDermott Technology
me: http://www.geocities.com/capecanaveral/3722 Please pardon the commercials!
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In a message dated 98-09-16 20:01:57 EDT, you write:
{ { Hi, I have a request for a micrograph of the pits and lands on a music CD. I thought this would be fun and simple but we aren't having any luck imaging anything (actually it was so reflective we imaged the detector!:-) Here's what we've done: We cut up a CD - using a piece of it close to the center hole (so we knew it had something on it to see). The piece was mounted (lower surface up) on
a stub using a carbon sticky tab and silver paste. The sample was sputter coated for 2 minutes (some for 3) and scoped. The first time we viewed it we didn't coat it (see paragraph one for details).
Do we need a solvent to munch awhile on the surface layer of the CD? Any help would be greatly appreciated!
It was a Christmas music CD so maybe that is the problem :-).
beth } }
How about making a replica and imaging it in the TEM?
If you wish to do that I can give you some technique hints.
Ted, I have good luck with CD images. Gold coat then tilt your sample as much as you can in your chamber, up to 85 degrees if possible. You should be able to see the pits and any defects such as shoulders around the pits, if present. I got a circular section (about 1.5 inch diameter) of the CD without introducing any stress by having it punched out in the machine shop. Hope this helps. ---------- } From: DUNNTEM-at-aol.com To: beth; Jackie Terry; Wayne England Cc: Microscopy -----------------------------------------------------------------------.
In a message dated 98-09-16 20:01:57 EDT, you write:
{ { Hi, I have a request for a micrograph of the pits and lands on a music CD. I thought this would be fun and simple but we aren't having any luck imaging anything (actually it was so reflective we imaged the detector!:-) Here's what we've done: We cut up a CD - using a piece of it close to the center hole (so we knew it had something on it to see). The piece was mounted (lower surface up) on a stub using a carbon sticky tab and silver paste. The sample was sputter coated for 2 minutes (some for 3) and scoped. The first time we viewed it we didn't coat it (see paragraph one for details).
Do we need a solvent to munch awhile on the surface layer of the CD? Any help would be greatly appreciated!
It was a Christmas music CD so maybe that is the problem :-).
beth } }
How about making a replica and imaging it in the TEM?
If you wish to do that I can give you some technique hints.
On Wed, 16 Sep 1998 DUNNTEM-at-aol.com-at-sparc5.microscopy.com wrote:
} I have a request for a micrograph of the pits and lands on a music CD. I } thought this would be fun and simple but we aren't having any luck imaging } anything (actually it was so reflective we imaged the detector!:-)
The pits and lands on the CD are on a metal layer that is sandwiched with a plastic one. The best way to image the metal surface is to delaminate the CD.
Nan Laudenslager wrote: ================================================ I have been asked to compare the grain size of 2 MgO aggregates. The samples were embedded and polished with diamond paste, but I am not happy with the result. The surface looks smeared. Does the sample need to be etched or do I need to evaluate my polishing technique???? ================================================= Certainly if there is room for improvement in the polishing technique, that is the first thing to do. However, when all else fails and you still have smearing (not unusual), then the smeared polymer can be selectively removed using a barrel type (isotropic) RF plasma etcher, using of course, oxygen. It is quite effective and it is a room temperature process. It takes only a few minutes of etching if that much.
Several commercially available table top etchers are available including our SPI Plasma Prep II, the details of which are on our website 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 seldom evaporate Au, but use W-wire baskets when I do... My source of Au is from "spent" sputter targets from years past. Why buy wire when I have a few perforated sputter targets around!
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I used to evaporate Au in a tungsten basket in an old Denton evap coater. We simply took a length of Au wire, wadded it up into a tiny ball and placed that in the basket. Took no large amount of manual dexterity and gave good results.
Gerroir, Paul J wrote:
} Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
HELP! I am new to the microscopy listserver and I am currently conducting undergraduate research for the Department of Biomedical Sciences at Southwest Missouri State University, located in Springfield, Missouri (USA). I am attempting to develop a TEM protocol to study the gap junction protein, Connexin-43 (Cx43) at the ultrastructural level. I am specifically interested in a protocol that utilizes colloidal gold as a marker and a post embedding technique that uses conventional resins. If anyone would be willing to share a specimen protocol for the localization of Cx43 or any other Connexins, please reply to me off list at: aaronrea-at-hotmail.com Thank you very much for your time!
Aaron Rea Department of Biomedical Sciences Southwest Missouri State University aaronrea-at-hotmail.com
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The simplest way that was suggested by Brian McIntyre and what worked fine was to simply scratch the back surface with a knife and take some good scotch tape and press on the back over the scratch with the tape. When it has been pressed onto the back with a lot of pressure, simply peel it off. You'll see the pattern transferred to the tape. Since this layer is conductive, there is no need to coat it as long as you have a good conductive path from that surface to the support stub. You can use carbon paint.
If you have the conductive carbon double sticky pads, they work well also.
-Scott Walck Scott D. Walck, Ph.D. PPG Industries, Inc. 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."
---------- } From: beth-at-dogwood.botany.uga.edu To: Microscopy-at-sparc5.microscopy.com
Hi, I have a request for a micrograph of the pits and lands on a music CD. I thought this would be fun and simple but we aren't having any luck imaging anything (actually it was so reflective we imaged the detector!:-) Here's what we've done: We cut up a CD - using a piece of it close to the center hole (so we knew it had something on it to see). The piece was mounted (lower surface up) on a stub using a carbon sticky tab and silver paste. The sample was sputter coated for 2 minutes (some for 3) and scoped. The first time we viewed it we didn't coat it (see paragraph one for details).
Do we need a solvent to munch awhile on the surface layer of the CD? Any help would be greatly appreciated!
It was a Christmas music CD so maybe that is the problem :-).
beth
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
We have several old American Optical/Spenser stereo microscopes that need a round glass stage insert. The inserts are 4" (10 cm) in diameter. I know we can have these made locally by a glass shop but was wondering whether anyone knows of an "off the shelf" source.
I am looking for the source of microparticles to colibrate/model light absorption processes. It can be latex spheres with diameter 1-10 um, partly stained with dyes(is it possible?). Any input will be appreciated.
Thanks
Dmitri Lapotko Luikov Heat and Mass Transfer Institute Minsk Belarus tel:(375172)842483 e-mail:ld-at-ns1.hmti.ac.by
Yes, having TWO (or more) tungsten filaments twisted together is important, to give the capillary action for more even dispersal as you mention. I forgot that important detail (it's been 27 years!) Thanks for jogging my memory. In fact, as I remember now, we were actually taking a long strand of tungsten wire and doubling it over more than once, to give a twisted set of 4 or more for our apparatus. We then wrapped the gold wire around that.
Regs, -- Tim
Luc Harmsen wrote: } Hi all. } Great topic. } We have been involved in building a new control unit for a factory which evaporates aluminium onto plastic parts to give them that silver look. } This has given us a lot more understanding of the coating technique as they have very large vacuum chambers and many samples that need an even and smooth coat each time. } What they do is to use tungsten wires coils that have two strands of W wire twisted together as aposed to, what we all seem to use, the single strand of W wire. The aluminium wire is then twisted by hand onto this coil, fairly loosely but just that it makes contact. The reasoning is that as you start the heating of the aluminium it will melt and flow, via capillary action, between the two W wires. This means that the aluminium is then in very good contact with the W coil and a lower current is needed to vaporise the aluminium. You also have a source of aluminium the whole length of the W coil each time. This ensures a even and repeatable coat each time. } } Cheers } Luc Harmsen } Anaspec, South Africa } International technical support on microscopy. } Tel: +27 (0) 11 476 3455 } Fax:+27 (0) 11 476 7290 } anaspec-at-icon.co.za } } -----Original Message----- } } From: Timothy Moeller [SMTP:tmoeller-at-noran.com] } Sent: Wednesday, September 16, 1998 7:15 PM } To: Paul.Gerroir-at-crt.xerox.com } Cc: Microscopy-at-sparc5.microscopy.com; Microscopy-at-sparc5.microscopy.com } Subject: Re: Evaporation of Gold } } Hi, Paul. } } Well, you've probably heard enough ideas on this subject. But I'll give } you my $0.02 worth (that's Au standard :-).) } } Working as a lab technician at UW Astrophysics Lab in Madison, WI, I } operated a vacuum deposition apparatus invented by a grad student which } vaporized gold wire and deposited it on the surface of insulated copper } wire to form the anode of a proportional detector. Anyway, in this } apparatus the gold wire was simply wrapped around tungsten (heater } filament) wire by hand. In vacuum, the tungsten filament was heated and } the gold wire melted, resulting in gold vapor being deposited evenly on } the wire rotissing nearby in the apparatus. It all seemed pretty crude } to me, but worked just fine. And I didn't take any special precautions } in wrapping the gold wire around the tungsten heater wire. } } Regs, } -- Tim } --- } } Gerroir, Paul J wrote: } } Hello Interested Readers, } } } } I have some experience in evaporating silver, (2 - 3mm shot) from a } } tungsten wire basket, however, I am now faced with challenge of evaporating } } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } } wire around the larger diameter tungsten wire and proceed or is there a } } better approach. Your comments/suggestions are appreciated. Thanks. } } } } Regards, } } Paul Gerroir } } Xerox Research Center of Canada }
-- ..we now return control of your computer screen to you... ---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ---------------------------------------------------------- "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ----------------------------------------------------------
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Hi, Paul.
Well, you've probably heard enough ideas on this subject. But I'll give you my $0.02 worth (that's Au standard :-).)
Working as a lab technician at UW Astrophysics Lab in Madison, WI, I operated a vacuum deposition apparatus invented by a grad student which vaporized gold wire and deposited it on the surface of insulated copper wire to form the anode of a proportional detector. Anyway, in this apparatus the gold wire was simply wrapped around tungsten (heater filament) wire by hand. In vacuum, the tungsten filament was heated and the gold wire melted, resulting in gold vapor being deposited evenly on the wire rotissing nearby in the apparatus. It all seemed pretty crude to me, but worked just fine. And I didn't take any special precautions in wrapping the gold wire around the tungsten heater wire.
Regs, -- Tim ---
Gerroir, Paul J wrote: } Hello Interested Readers, } } I have some experience in evaporating silver, (2 - 3mm shot) from a } tungsten wire basket, however, I am now faced with challenge of evaporating } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } wire around the larger diameter tungsten wire and proceed or is there a } better approach. Your comments/suggestions are appreciated. Thanks. } } Regards, } Paul Gerroir } Xerox Research Center of Canada
-- ...we now return control of your computer screen to you... ------------------------------------------------------------ Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments, Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ------------------------------------------------------------ "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ------------------------------------------------------------
SCANNING PROBE MICROSCOPY: PRINCIPLES AND PRACTICE.
18-22 January 1999
Short Course from the School of Mechanical and Materials Engineering, University of Surrey, Guildford, UK.
THE COURSE
The aim of this five day intensive course is to introduce the principles and practice of Scanning Tunnelling Microscopy (STM), Scanning Force Microscopy (SFM) and other methods of Scanning Probe Microscopy (SPM). The physical concepts employed in the instrumentation of STM and SFM are simple, but the interpretation of the STM and SFM results can be complicated because of the convolution of several interactions in the measurement process. This complication exists in the large scale imaging of surface morphology as well as in the molecular- and atomic scale images. Thus, many STM and SFM studies can be misinterpreted. To help to alleviate this problem, we felt it necessary to bring together in this course the essential components of STM and SFM studies, namely, the practical aspects of STM and SFM, the image simulation and the qualitative evaluations of tip force induced surface corrugations.
The primary goal of the course will be to describe how the surfaces of various materials are characterised by employing STM, SFM and other methods of SPM and what physical/chemical features can be deducted from their images.
This will be achieved through a balance of lectures, tutorials and laboratory demonstrations. The course will provide a theoretical introduction to the field and an overview of recent development. Lectures given by leading SPM experts will be supported by supervised exercise classes in which experience will be gained in the solution of typical problems in SPM.
The Course will cover the basics of operation and advanced operation. Course registrants will have access to two STM and SFM microscopes in the Surface and Interface Reaction Group and much of the subject matter will be demonstrated on these instruments.
WHO SHOULD ATTEND
The course will be of maximum benefit to you if you are, or expect to be involved in using any form of Scanning Probe Microscopy as a research, diagnostic or trouble-shooting tool. Engineers, Chemists and Physicists who are using Scanning Electron Microscopy and Transmission Electron Microscopy will also find the course extremely useful.
COURSE FORMAT
The course will commence with registration at 9:30 on Monday 18 January 1999 and continue until 15:00 on Friday 22 January. The program of lectures is well distributed with a variety of tutorials each day. There will be plenty of opportunities for discussion with lecturers and other delegates. Full course notes will be supplied to all participants.
ORGANISERS AND PRINCIPAL LECTURERS:
Professor Jim Castle (SMME, University of Surrey) Dr. Peter Zhdan (SMME, University of Surrey)
INVITED LECTURERS INCLUDE:
Professor Michael Bowker (University of Reading) Professor Martyn Davies (University of Nottingham) Professor Trevor Page (University of Newcastle) Professor John Pethica (University of Oxford) Professor Richard Palmer (University of Birmingham)
For further information contact Dr. Peter Zhdan (P.Zhdan-at-surrey.ac.uk) or the Course Secretary:
Mrs. Margaret Morgan School of Mechanical and Materials Engineering University of Surrey Guildford, Surrey GU2 5XH UK
} -----Original Message----- } From: Steve Chapman [mailto:PROTRAIN-at-CompuServe.COM] } Sent: Thursday, September 17, 1998 12:04 AM } To: Melvyn Dickson } Cc: [unknown] Mel had written ... } } } } His answer was that users were opening the inner door of the airlock too } } soon and perhaps too quickly. There was too much air left in the airlock } } and the pressure rise in the chamber stalled the diffusion pump and } } backstreaming of the diffusion pump oil caused the contamination. }
Steve responded ... } } Do you believe this guy? } } All the work we did in the past proved to us beyond doubt } that the oil was from the rotary pump. No matter how } sophisticated the system is if you use } a rotary pump at all in the cycle you do seem to get RP oil as a } contaminant. } } I think the guy had a good try but I for one do not believe him! } } ...
I believe what the JEOL rep suggested was not out of line ... the contamination would still be from the RP if there were a slight DP burp.
What many JEOL users are not aware of is that (1) the "ready" lite responds to a timer not the interlock vacuum and (2) since JEOL started relying on a single RP (i.e., an RP is no longer 100% dedicated to the DP), the interlock pressure will rise after "ready" is indicated. I have to make sure all my users know this and that they should exchange the specimen immediately. Also (3) the interlock seal is subject to leaks because of dust, therefore the "ready" lite should be considered an indication of 90seconds only ... it is not an indicator that the interlock vacuum is ready. Given these possible problems, I wish JEOL would cap the DP for specimen exchanges ...
cheerios, shAf
{} /\ {\/} /\ {\/} /\ {\/} /\ cogito, ergo zZOooOM /\ {\/} /\ {\/} /\ {\/} /\ {} Michael Shaffer, R.A. - ICQ 210524 Geological Science's Electron Probe Facility - University of Oregon mshaf-at-darkwing.uoregon.edu - http://darkwing.uoregon.edu/~mshaf/
What are the lateral and Z dimensions of these structures? There have been a number of other techniques,including SWLI (Scanning White Light Interferometry) used to image these structures. I have access to a system which can do that, if it would be helpful.
I didn't think there was this much gold in the world to evaporate!
****************************** Jim Haley Applications Engineer I-CUBE 2411 Crofton Lane, Suite 14A Crofton, MD 21114 voice: (301) 858-0505 fax: (301) 858-0615 web site: http://www.i-cubeinc.com e-mail: haley-at-i-cubeinc.com ******************************
Timothy Moeller 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 } -----------------------------------------------------------------------. } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } ------------------------------------------------------------------------ } The 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, Paul. } } Well, you've probably heard enough ideas on this subject. But I'll give } you my $0.02 worth (that's Au standard :-).) } } Working as a lab technician at UW Astrophysics Lab in Madison, WI, I } operated a vacuum deposition apparatus invented by a grad student which } vaporized gold wire and deposited it on the surface of insulated copper } wire to form the anode of a proportional detector. Anyway, in this } apparatus the gold wire was simply wrapped around tungsten (heater } filament) wire by hand. In vacuum, the tungsten filament was heated and } the gold wire melted, resulting in gold vapor being deposited evenly on } the wire rotissing nearby in the apparatus. It all seemed pretty crude } to me, but worked just fine. And I didn't take any special precautions } in wrapping the gold wire around the tungsten heater wire. } } Regs, } -- Tim } --- } } Gerroir, Paul J wrote: } } Hello Interested Readers, } } } } I have some experience in evaporating silver, (2 - 3mm shot) from a } } tungsten wire basket, however, I am now faced with challenge of evaporating } } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } } wire around the larger diameter tungsten wire and proceed or is there a } } better approach. Your comments/suggestions are appreciated. Thanks. } } } } Regards, } } Paul Gerroir } } Xerox Research Center of Canada } } -- } ...we now return control of your computer screen to you... } ------------------------------------------------------------ } Timothy G. Moeller | Microanalysis Products } Senior Software Engineer | NORAN Instruments, Inc., } {tmoeller-at-noran.com} | a ThermoSpectra company } ------------------------------------------------------------ } "I've spent my whole life trying to think up crazy ways of } doing things." } - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") } ------------------------------------------------------------
I am working with pollen and anther development of Ilex paraguariensis St. Hil. (Aquifoliaceae). However, the techniques used by me does not result in good ultrastrutural preservation. I am using Glutaraldehyde 2% + paraformaldehyde 2% in phosphate buffer ph 7.2, 0.1 M, at 4 oC, as a primary fixative. How obtain a "live-like" strutures in TEM? Which fixative should be used (concentrations, pH, osmolarity...)?
Thanks in advance M.Sc. Rinaldo Pires dos Santos Dept. of Botany - Universidade Federal do Rio Grande do Sul - UFRGS Av. Paulo Gama, 40 - Bairro Bom Fim - 90046-900 Porto Alegre - RS - Brazil e-mail: rinaldop-at-botanica.ufrgs.br
Dear Mel, This is a problem of all SEMs, not just JEOL, and really is a result of the EDS detector being the coolest spot in the chamber. The cleanliness of the vacuum system just regulates how long the contamination will take to build up. Link solved this problem by warming the snout of their detector. This doesn't solve the problem of oil contamination, it just moves it away from the EDS detector. I agree woth Steve Chapman that the oil is from the rotary pump. You wrote:
} Numbers of JEOL owners in Oz have complained of similar oil contamination. } When we last evaluated FESEMs the JEOL rep was persistent to know why we } didn't like the JEOL models. One of my reasons was the oil contamination } problem and I asked for some explanation as to why it was so common. } } His answer was that users were opening the inner door of the airlock too } soon and perhaps too quickly. There was too much air left in the airlock } and the pressure rise in the chamber stalled the diffusion pump and } backstreaming of the diffusion pump oil caused the contamination. } } } ***************************************************** } Mel Dickson, 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 fax: 604-822-3619 e-mail: mager-at-interchg.ubc.ca
Dear Beth, I have prepared CDs for SEM imaging and you must dissolve the plastic to get at the very thin aluminum foil that has the pits on it. The laser reads the pits through the plastic. I used tri-chlorethylene overnight to dissolve the plastic from a 1 square cm piece of the outside edge of a CD. In the morning I fished out this tiny scrap of Al foil and put it on an SEM stub. You can image either side and at 10,000 times see the little pits in their rows. These are pre-recorded CDs, the CD-Rs are quite different, since they use a laser-sensitive dye. You wrote: } } Hi, } I have a request for a micrograph of the pits and lands on a music CD. I } thought this would be fun and simple but we aren't having any luck imaging } anything (actually it was so reflective we imaged the detector!:-) } Here's what we've done: } We cut up a CD - using a piece of it close to the center hole (so we knew } it had something on it to see). The piece was mounted (lower surface up) on } a stub using a carbon sticky tab and silver paste. The sample was sputter } coated for 2 minutes (some for 3) and scoped. The first time we viewed it } we didn't coat it (see paragraph one for details). } } Do we need a solvent to munch awhile on the surface layer of the CD? } Any help would be greatly appreciated! } } It was a Christmas music CD so maybe that is the problem :-). } } beth Good luck, 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 fax: 604-822-3619 e-mail: mager-at-interchg.ubc.ca
At the risk of further crowding your in-basket(s), I want to apologize to everyone for my recent mishap in posting an article (Re: Evaporation of Gold) which resulted in bouncing ad infinitum (or ad nauseum, if you prefer.) I believe the problem stems to a mistake I made in replying to the list on that subject. Apparently, you are not supposed to use the "Reply" function, but rather post a NEW message to the List Server at {Microscopy-at-MSA.Microscopy.Com} . So consider this a warning in addition to an apology -- it could happen to you as easily as it has happened to me. In fact, I'd seen this same thing happen to other posters as well recently, who apparently made the same mistake I did.
---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ----------------------------------------------------------
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Hi, Luc et al.
Yes, having TWO (or more) tungsten filaments twisted together is important, to give the capillary action for more even dispersal as you mention. I forgot that important detail (it's been 27 years!) Thanks for jogging my memory. In fact, as I remember now, we were actually taking a long strand of tungsten wire and doubling it over more than once, to give a twisted set of 4 or more for our apparatus. We then wrapped the gold wire around that.
Regs, -- Tim
Luc Harmsen wrote: } Hi all. } Great topic. } We have been involved in building a new control unit for a factory which evaporates aluminium onto plastic parts to give them that silver look. } This has given us a lot more understanding of the coating technique as they have very large vacuum chambers and many samples that need an even and smooth coat each time. } What they do is to use tungsten wires coils that have two strands of W wire twisted together as aposed to, what we all seem to use, the single strand of W wire. The aluminium wire is then twisted by hand onto this coil, fairly loosely but just that it makes contact. The reasoning is that as you start the heating of the aluminium it will melt and flow, via capillary action, between the two W wires. This means that the aluminium is then in very good contact with the W coil and a lower current is needed to vaporise the aluminium. You also have a source of aluminium the whole length of the W coil each time. This ensures a even and repeatable coat each time. } } Cheers } Luc Harmsen } Anaspec, South Africa } International technical support on microscopy. } Tel: +27 (0) 11 476 3455 } Fax:+27 (0) 11 476 7290 } anaspec-at-icon.co.za } } -----Original Message----- } } From: Timothy Moeller [SMTP:tmoeller-at-noran.com] } Sent: Wednesday, September 16, 1998 7:15 PM } To: Paul.Gerroir-at-crt.xerox.com } Cc: Microscopy-at-sparc5.microscopy.com; Microscopy-at-sparc5.microscopy.com } Subject: Re: Evaporation of Gold } } Hi, Paul. } } Well, you've probably heard enough ideas on this subject. But I'll give } you my $0.02 worth (that's Au standard :-).) } } Working as a lab technician at UW Astrophysics Lab in Madison, WI, I } operated a vacuum deposition apparatus invented by a grad student which } vaporized gold wire and deposited it on the surface of insulated copper } wire to form the anode of a proportional detector. Anyway, in this } apparatus the gold wire was simply wrapped around tungsten (heater } filament) wire by hand. In vacuum, the tungsten filament was heated and } the gold wire melted, resulting in gold vapor being deposited evenly on } the wire rotissing nearby in the apparatus. It all seemed pretty crude } to me, but worked just fine. And I didn't take any special precautions } in wrapping the gold wire around the tungsten heater wire. } } Regs, } -- Tim } --- } } Gerroir, Paul J wrote: } } Hello Interested Readers, } } } } I have some experience in evaporating silver, (2 - 3mm shot) from a } } tungsten wire basket, however, I am now faced with challenge of evaporating } } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } } wire around the larger diameter tungsten wire and proceed or is there a } } better approach. Your comments/suggestions are appreciated. Thanks. } } } } Regards, } } Paul Gerroir } } Xerox Research Center of Canada }
-- .we now return control of your computer screen to you... ---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ---------------------------------------------------------- "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ----------------------------------------------------------
Arthropods generally have problems like this, but they're easily solved by dissection. The problem is the cuticle.
You need to create as good a path for the fluids to flow through the critters as possible: pull off a leg and snip off the tarsus if you want to look at leg structures, etc.
Remove the opisthosoma and a couple of legs if you're interested in the prosoma, etc.
Phil
} Greetings, } } I would like to fix small spiders for TEM. I am not particular interested } in preservation of specific structures, but would like to preserve both } integumental and inner structures in general. I noticed that my specimens } floated, and I am afraid that the fixation will not penetrate properly. } } Does anybody has experience with fixation of similar specimens ? I would } appreciate your comments very much. } } Thank you. } } Regards } } Peter Funch } } ______________________________________________________________________ } Peter Funch } Assistant Professor, Ph.D. } } Department of Zoology Direct Line + 45 8942 2764 } Institute of Biological Sciences Secretary + 45 8942 2727 } University of Aarhus Telefax + 45 86 12 51 75 } Universitetsparken E-mail: } peter.funch-at-biology.aau.dk } Building 135 } DK-8000 Aarhus C } Denmark } ______________________________________________________________________
}}}}}}}}}}}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{{{{{{{{{{ Philip Oshel PO Box 620068 Middleton, WI 53562 (608) 833-2885 oshel-at-terracom.net or poshel-at-hotmail.com
Does anyone know why some Polaroid shots from an SEM have faint parallel vertical lines with a consistant spacing of somewhat less than 0.5 mm? Are they in the film? I'm also getting patches of horizontal lines in some micrographs lately that I'm pretty sure aren't due to either charging or the Polaroid rollers. Could this be interference from the new lab next door?
Thanks,
Dee
____________________________________________________________________________ Note: Sometimes I don't receive incoming emails (with no notification to the sender). If I don't respond to your message, please send it again! ____________________________________________________________________________ _ Dee Breger Manager, SEM/EDX Facility Lamont-Doherty Earth Observatory Route 9W Palisades NY 10964 USA
To anyone who cares, After 50 hrs in 1% alconox the appearance and mechanical proper- ties of the Al foil seem to be unchanged. I didn't weigh the foil or look at it under a microscope, but anyone with too much free time is welcome to try it. Yours, Bill Tivol
I have some images of a CD master taken by SEM and AFM if you would like them. Drop me a line and I can send them as an attachement.
JB
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing 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 ####################################################################
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I am presently researching digital cameras which can be mounted on light microscopes for use for both fluorescence and bright field image capture. This is to augment film recording, not replace it. Resolution is very important as is even light spread. I am also interested in the different image storage mechanisms utilized by these cameras.
Any suggestions or personal experiences with using digital cameras for this purpose would be appreciated.
I promise to post a summary of responses.
Thanks, Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
} Does anyone know why some Polaroid shots from an SEM have faint parallel } vertical lines with a consistant spacing of somewhat less than 0.5 mm? Are } they in the film? I'm also getting patches of horizontal lines in some } micrographs lately that I'm pretty sure aren't due to either charging or } the Polaroid rollers. Could this be interference from the new lab next } door?
#################################################################### John J. Bozzola, Ph.D., Director Center for Electron Microscopy Neckers Building, Room 146 - B Wing 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 ####################################################################
We make up a saturated stock bottle which we draw from, and replenish with UA and water (8-10g UA/100 ml) from time to time. The insoluble material is described in the Merck Index as being due to insoluble basic salts. It describes Uranyl Acetate as being "freely soluble in water acidulated with acetic acid" For years, we have followed a modification of a procedure from Millonig's 1976 book Laboratory Manual of Biological Electron Microscopy (pg 53) and added a few drops of acetic acid per 100mls of stock saturated UA (stored in a brown bottle). This seems to push the ppt reaction the other way and give a clear solution. There seems to be little difference in staining as long as only a few drops of acetic acid are used. Changing the pH of the stain by much, is risky though as there are numerous papers and procedures which modify the effects of UA stain by doing so. We have raised the pH to the 4.5-5.5 (any higher and the UA will ppt) and gotten improved staining but with unacceptable amounts of ppt on the sections.
When compared with the other chemicals in the EM lab, UA would seem to be relatively safe when used carefully. Ingestion and inhalation (exposure to dust) are our major concern due to heavy metal toxicity as well as the radiation hazards. Making sure that surfaces are not contaminated, and cleaning any spillage immediately from bottles and tables before it dries are important steps. Wearing gloves, and hand washing after glove removal are also important safeguards.
The radiation exposure hazard under most operating conditions seems minimal. The least exposure possible is desirable (ALARA), when you don't need to handle it, don't be near it. Using Bill's number's, you would still be well under the limits for occupational exposure if you were in constant contact with .6 millirem/hr for a 2000 hr work year, (correct me, but my references place the limits at 1.25 rem/quarter, 5 rem/year whole body and 18.75 rem/quarter, 75 rem/year for extremities (Rayburn)) The other factor to keep in mind is that we are not talking about a whole body exposure, but just exposure to the hands. All in all, the amount of exposure while making up and staining grids seems miniscule.
As an aside, the pretty flowered dinnerware from the 50's, the vivid oranges and yellows are from uranium. If you have any, run a Geiger counter over them, you'll be surprised the number of counts. Also the mantles from gas and propane lanterns contain radioactive thorium. In the past health physicist have suggested using them(sealed in their bags) for check sources for counters.
Regardless, because of the toxicity, radiation hazard, as well as expenses to purchase(well over $1.00/gm) and dispose of UA, minimizing the amount needed to be discarded and wasted seems desirable. To the extents possible, use of minimal amounts, and if considerable staining is done, making stock saturated solutions which can be diluted to the desired concentration as needed, are good ways to conserve UA, minimize radiation exposure, and inhalation and ingestion hazards.
Now, if we are starting a poll for the chemicals in the EM Lab that make us the most anxious, my vote is for cacodylate.
Excellent. We are also in the process of researching digital cameras for the same purpose. Any words of experience would be greatly appreciated. Linda Barthel Research Associate II Department of Anatomy and Cell Biology University of Michigan lab (313) 764-7476 fax (313) 763-1166 barthel-at-umich.edu
On 17 Sep 1998, Debby Sherman 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 presently researching digital cameras which can be mounted on } light microscopes for use for both fluorescence and bright field image } capture. This is to augment film recording, not replace it. Resolution is very } important as is even light spread. I am also interested in the different } image storage mechanisms utilized by these cameras. } } Any suggestions or personal experiences with using digital cameras for } this purpose would be appreciated. } } I promise to post a summary of responses. } } Thanks, } Debby Sherman, Manager Phone: 765-494-6666 } Microscopy Center in Agriculture FAX: 765-494-5896 } Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu } Purdue University } 1057 Whistler Building } West Lafayette, IN 47907-1057 } }
Beth- here is the technique I developed at UW (Seattle) with the help of a few of the gradstudents, it worked for me, hope it works for you. -Mike
Protocol for preparation of CDs for SEM analysis (by Mike Rock)
This simple method utilizes the coefficient of thermal expansion for separation of materials of differing densities. CDs are made up of a metallic core (usually aluminum or gold) surrounded by a plastic layer on either side. Other methods include dissolving the plastic with various solvents, or by removing the metal layer by etching techniques. Both may work fine, I have tried neither. This protocol uses liquid nitrogen to cool the sample (CD) to a point where the materials separate, and has proved successful with both gold and aluminum CDs.
Using tongues immerse the CD in the liquid nitrogen, after 15-30 seconds the CD will sound as if it is cracking. After 30- 60 seconds remove the CD from the liquid nitrogen.
Place the frozen CD on a firm surface and strike it with a hammer (wear safety glasses), the CD will shatter. Alternatively you may wish to slap the frozen CD down against the bench top (results of the two techniques are similar), shearing between the plastic and metal interface. The metal will easily pull away from the surface of the plastic if still in contact.
Mount the metallic layer, which contains the information tracks ("pits" and "lands") on a aluminum stub using double stick "conductive" carbon tape or tabs. Sputter coating is usually not necessary. Examination with the SEM is fairly routine at this point (5-15 kV).
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Hi, Luc et al.
Yes, having TWO (or more) tungsten filaments twisted together is important, to give the capillary action for more even dispersal as you mention. I forgot that important detail (it's been 27 years!) Thanks for jogging my memory. In fact, as I remember now, we were actually taking a long strand of tungsten wire and doubling it over more than once, to give a twisted set of 4 or more for our apparatus. We then wrapped the gold wire around that.
Regs, -- Tim
Luc Harmsen wrote: } Hi all. } Great topic. } We have been involved in building a new control unit for a factory which evaporates aluminium onto plastic parts to give them that silver look. } This has given us a lot more understanding of the coating technique as they have very large vacuum chambers and many samples that need an even and smooth coat each time. } What they do is to use tungsten wires coils that have two strands of W wire twisted together as aposed to, what we all seem to use, the single strand of W wire. The aluminium wire is then twisted by hand onto this coil, fairly loosely but just that it makes contact. The reasoning is that as you start the heating of the aluminium it will melt and flow, via capillary action, between the two W wires. This means that the aluminium is then in very good contact with the W coil and a lower current is needed to vaporise the aluminium. You also have a source of aluminium the whole length of the W coil each time. This ensures a even and repeatable coat each time. } } Cheers } Luc Harmsen } Anaspec, South Africa } International technical support on microscopy. } Tel: +27 (0) 11 476 3455 } Fax:+27 (0) 11 476 7290 } anaspec-at-icon.co.za } } -----Original Message----- } } From: Timothy Moeller [SMTP:tmoeller-at-noran.com] } Sent: Wednesday, September 16, 1998 7:15 PM } To: Paul.Gerroir-at-crt.xerox.com } Cc: Microscopy-at-sparc5.microscopy.com; Microscopy-at-sparc5.microscopy.com } Subject: Re: Evaporation of Gold } } Hi, Paul. } } Well, you've probably heard enough ideas on this subject. But I'll give } you my $0.02 worth (that's Au standard :-).) } } Working as a lab technician at UW Astrophysics Lab in Madison, WI, I } operated a vacuum deposition apparatus invented by a grad student which } vaporized gold wire and deposited it on the surface of insulated copper } wire to form the anode of a proportional detector. Anyway, in this } apparatus the gold wire was simply wrapped around tungsten (heater } filament) wire by hand. In vacuum, the tungsten filament was heated and } the gold wire melted, resulting in gold vapor being deposited evenly on } the wire rotissing nearby in the apparatus. It all seemed pretty crude } to me, but worked just fine. And I didn't take any special precautions } in wrapping the gold wire around the tungsten heater wire. } } Regs, } -- Tim } --- } } Gerroir, Paul J wrote: } } Hello Interested Readers, } } } } I have some experience in evaporating silver, (2 - 3mm shot) from a } } tungsten wire basket, however, I am now faced with challenge of evaporating } } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } } wire around the larger diameter tungsten wire and proceed or is there a } } better approach. Your comments/suggestions are appreciated. Thanks. } } } } Regards, } } Paul Gerroir } } Xerox Research Center of Canada }
-- .we now return control of your computer screen to you... ---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ---------------------------------------------------------- "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ----------------------------------------------------------
This bounced once, I shall try again... (Nestor ignore the (not spam) email if this makes it to the listserver ok) ----------------------------------------------------------------
Well....
The Nuclear Regulatory Commission limits of skin & extremity (hands, feet) is 50 Rem per year. ...Not something to "shoot" for, since the dose is also limited to "As Low As Reasonably Achievable".
The (damage) conversion coefficient from mR from this source (no alpha if not ingested) to mRem is ~1. 50 Rem = 50,000 mRem. At a dose rate of 0.6 mR/hr, one would have to hold the container for many years to receive a one year maximum dose (50,000 / 0.6 per hr = max hours exposure). At 5 mR/hr, it would be 50,000 / 5. At that one would have to hold the container for 10,000 hours before exceeding NRC dose limits. Exposure will also decrease as a function of the square of the distance from the source.
For medical tests to discover any changes in body chemistry, it would take about 50 Rem acute whole body exposure.
Less dose is always better, but in realistic terms the dose from the UA should not be of any concern. If this level is of concern, do not fly in airplanes, live at high elevations, avoid all medical radiation, avoid certain beaches, beware of granite buildings, run from radium dial watches, etc. :)
The real danger is if the UA enters the body where the alpha source is in direct contact with livings tissue. Radiological bio-assay (urine/fecal) would be required to detect this.
} Using Bill's number's, you would } still be well under the limits for occupational exposure if you were in } constant contact with .6 millirem/hr for a 2000 hr work year, (correct me, } but my references place the limits at 1.25 rem/quarter, 5 rem/year whole } body
These limits are for radiation workers. Because we get paid, we can be exposed to a greater risk. The limit for the general population is 0.5 rem/year whole body, and I think this limit also applies to women who are or may be pregnant. I do not know the status of graduate students; I'd be inclined to err on the side of caution--especially since it is fairly easy to keep exposure to UA low. Yours, Bill Tivol
Look, I'm sorry. But this wasn't my doing. True, it stems from an oversight on my part (by posting to the wrong address, as I explained in my apology article), but I did NOT compound the error myself by re-sending the message, nor repeating the CC's, nor anything. The listserver did all that. This whole thing just blew up in my face, after I posted one simple little article incorrectly. I'm sorry, but it's out of my control.
Nestor -- please fix this at the server!
Everyone else -- please try to understand, I'm sorry!
You can improve membrane fixation in a couple of ways. The best I use is pos-fixation in 1% Uranyl acetate in bidestilled water = for 1hour (do not use Cacodilate or Phosphate buffers since the uranyl = acetate precipitates). This is done after the Osmium tetroxide fixation, = so, be carefull to wash these buffers out of the material before = applying the Uranyl acetate.=20 You can use if you wish 0.1M acetate acetic acid buffer. The pH should = be lower than 6.7 (? must check this value) since Uranyl acetate = precipitates at higher pH.
You can also add Potassium ferricyanide 0.5% to the osmium tetroxide and = fix longer than usual (up to 5 hours).
You may also have problems with the embeding medium. Some embeddings may = produce bad membrane preservation. I use Molenhauer's EPON-ARALDITE = mixture with good results.
Hope this helps
Dr. A.P. Alves de Matos Pathology Department Curry Cabral Hospital Lisbon apmatos-at-ip.pt
{!DOCTYPE HTML PUBLIC "-//W3C//DTD W3 HTML//EN"} {HTML} {HEAD}
{META content=3Dtext/html;charset=3Diso-8859-1 = http-equiv=3DContent-Type} {META content=3D'"MSHTML 4.72.3110.7"' name=3DGENERATOR} {/HEAD} {BODY bgColor=3D#ffffff} {DIV} {FONT color=3D#000000 size=3D2} You can improve membrane fixation in = a couple of=20 ways. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} The best I use is pos-fixation in 1% = Uranyl=20 acetate in bidestilled water for 1hour (do not use Cacodilate or = Phosphate=20 buffers since the uranyl acetate precipitates). This is done after the = Osmium=20 tetroxide fixation, so, be carefull to wash these buffers out of the = material=20 before applying the Uranyl acetate. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} You can use if you wish 0.1M acetate = acetic acid=20 buffer. The pH should be lower than 6.7 (? must check this value) since = Uranyl=20 acetate precipitates at higher pH. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} You can also add Potassium = ferricyanide 0.5% to=20 the osmium tetroxide and fix longer than usual (up to 5 = hours). {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} You may also have problems with the = embeding=20 medium. Some embeddings may produce bad membrane preservation. I use=20 Molenhauer's EPON-ARALDITE mixture with good results. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {/DIV} {DIV} {FONT size=3D2} Hope this helps {/FONT} {/DIV} {DIV} {FONT size=3D2} {/FONT} {/DIV} {DIV} {FONT size=3D2} Dr. A.P. Alves de Matos {/FONT} {/DIV} {DIV} {FONT size=3D2} Pathology Department {/FONT} {/DIV} {DIV} {FONT size=3D2} Curry Cabral Hospital {/FONT} {/DIV} {DIV} {FONT size=3D2} Lisbon {/FONT} {/DIV} {DIV} {FONT size=3D2} {A = href=3D"mailto:apmatos-at-ip.pt"} apmatos-at-ip.pt {/A} {/FONT} {/DIV} {DIV} {FONT size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {/BODY} {/HTML}
Group We hope to present the result of our salary survey in around a month. The hold up was caused by the lack of interest - particularly from this listserver. It has been, however, substantial from the readers of our publication and the MSA Conference attendees. If you are interested in the survey results, and have not yet provided your data (in absolute confidence), you are invited to do so. The survey includes only microscopists in the U.S. and excludes manufacturers and suppliers. Reponses may be made by return email or by fax (608-836-1969). As follows, your data should be in 8 fields: FIELD 1: Last Education Degree None/AA/BS/MS/PHD/MD FIELD 2: Years experience after graduation FIELD 3: Gender M - Male F - Female FIELD 4:Yearly income FIELD 5: Are you currently a supervisor/manager? Y - Yes N - No FIELD 6: Location. If in question, pick the area you feel closest to your own income level. MW - Midwest NE - Northeast SE- Southeast S - South W - West excluding California CA - California FIELD 7: Primary interest in: B - Biological Science P - Physical Science E - Earth Science FIELD 8: Now working in: I - Industry E - Education H - Hospital/Medical G - Government (as employee with GS Scale) GS - Government sponsered research
Immersion oil. Students do not always clean the oil off of the objectives after use, and thus we have suffered oil infiltration into some objectives. Recently one of our faculty members was advised to use type B immersion oil as opposed to type A oil. This due to the type objectives we are using, A.O. 100x oil and Olympus 100x oil. My questions, is there a chemical make up difference in these oils, or is it just a high-low viscosity issue. TIA tom_osborn-at-csubak.edu Tom Osborn Staff California State University Bakersfield
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Hi, Luc et al.
Yes, having TWO (or more) tungsten filaments twisted together is important, to give the capillary action for more even dispersal as you mention. I forgot that important detail (it's been 27 years!) Thanks for jogging my memory. In fact, as I remember now, we were actually taking a long strand of tungsten wire and doubling it over more than once, to give a twisted set of 4 or more for our apparatus. We then wrapped the gold wire around that.
Regs, -- Tim
Luc Harmsen wrote: } Hi all. } Great topic. } We have been involved in building a new control unit for a factory which evaporates aluminium onto plastic parts to give them that silver look. } This has given us a lot more understanding of the coating technique as they have very large vacuum chambers and many samples that need an even and smooth coat each time. } What they do is to use tungsten wires coils that have two strands of W wire twisted together as aposed to, what we all seem to use, the single strand of W wire. The aluminium wire is then twisted by hand onto this coil, fairly loosely but just that it makes contact. The reasoning is that as you start the heating of the aluminium it will melt and flow, via capillary action, between the two W wires. This means that the aluminium is then in very good contact with the W coil and a lower current is needed to vaporise the aluminium. You also have a source of aluminium the whole length of the W coil each time. This ensures a even and repeatable coat each time. } } Cheers } Luc Harmsen } Anaspec, South Africa } International technical support on microscopy. } Tel: +27 (0) 11 476 3455 } Fax:+27 (0) 11 476 7290 } anaspec-at-icon.co.za } } -----Original Message----- } } From: Timothy Moeller [SMTP:tmoeller-at-noran.com] } Sent: Wednesday, September 16, 1998 7:15 PM } To: Paul.Gerroir-at-crt.xerox.com } Cc: Microscopy-at-sparc5.microscopy.com; Microscopy-at-sparc5.microscopy.com } Subject: Re: Evaporation of Gold } } Hi, Paul. } } Well, you've probably heard enough ideas on this subject. But I'll give } you my $0.02 worth (that's Au standard :-).) } } Working as a lab technician at UW Astrophysics Lab in Madison, WI, I } operated a vacuum deposition apparatus invented by a grad student which } vaporized gold wire and deposited it on the surface of insulated copper } wire to form the anode of a proportional detector. Anyway, in this } apparatus the gold wire was simply wrapped around tungsten (heater } filament) wire by hand. In vacuum, the tungsten filament was heated and } the gold wire melted, resulting in gold vapor being deposited evenly on } the wire rotissing nearby in the apparatus. It all seemed pretty crude } to me, but worked just fine. And I didn't take any special precautions } in wrapping the gold wire around the tungsten heater wire. } } Regs, } -- Tim } --- } } Gerroir, Paul J wrote: } } Hello Interested Readers, } } } } I have some experience in evaporating silver, (2 - 3mm shot) from a } } tungsten wire basket, however, I am now faced with challenge of evaporating } } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } } wire around the larger diameter tungsten wire and proceed or is there a } } better approach. Your comments/suggestions are appreciated. Thanks. } } } } Regards, } } Paul Gerroir } } Xerox Research Center of Canada }
-- .we now return control of your computer screen to you... ---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ---------------------------------------------------------- "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ----------------------------------------------------------
You probably have to cut the animals in a drop of fixative or inject the = fixative into the animal's body. If the animal is too small, I would try to injure the surface to provide = entry points to the fixative. Perhaps removing the legs?. The floating can be controled by pushing the animals into the fixative = bottle with a cotton plug, filter paper or something similar.
Dr. A.P. Alves de Matos Pathology Department Curry Cabral Hospital Lisbon apmatos-at-ip.pt
{!DOCTYPE HTML PUBLIC "-//W3C//DTD W3 HTML//EN"} {HTML} {HEAD}
{META content=3Dtext/html;charset=3Diso-8859-1 = http-equiv=3DContent-Type} {META content=3D'"MSHTML 4.72.3110.7"' name=3DGENERATOR} {/HEAD} {BODY bgColor=3D#ffffff} {DIV} {FONT color=3D#000000 size=3D2} You probably have to cut the animals = in a drop=20 of fixative or inject the fixative into the animal's body. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} If the animal is too small, I would = try to=20 injure the surface to provide entry points to the fixative. Perhaps = removing the=20 legs?. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} The floating can be controled by = pushing the=20 animals into the fixative bottle with a cotton plug, filter paper or = something=20 similar. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} Dr. A.P. Alves de Matos {/FONT} {/DIV} {DIV} {FONT size=3D2} Pathology Department {/FONT} {/DIV} {DIV} {FONT size=3D2} Curry Cabral Hospital {/FONT} {/DIV} {DIV} {FONT size=3D2} Lisbon {/FONT} {/DIV} {DIV} {FONT size=3D2} {A=20 href=3D"mailto:apmatos-at-ip.pt"} apmatos-at-ip.pt {/A} {/FONT} {/DIV} {/BODY} {/HTML= }
Interesting - I have always hit the 'reply' button, intending to send something just to the individual doing the posting, and that is exactly what has happened, and there has been nothing at all on the ListServer (never mind 'bouncing ad infinitum'). I have just now hit the 'reply all' button on our MS Exchange system for this sending (with the difference that the 'Microscopy-at-Sparc5.Microscopy.Com' address is there along with yours). So let's see what transpires, but I have a feeling the 'bouncing' has something subtle to do with your local stuff. I'm sure our esteemed Sysop would have posted 'How To Avoid' guidelines by now if it was as simple a matter as you say. (Or----, Nestor?).
tom
Tom Malis Group Leader - Characterization Materials Technology Laboratory Natural Resources Canada (Govt. of Canada) 568 Booth St., Ottawa, Canada ph. 613-992-2310 FAX 623-992-8735 email: malis-at-nrcan.gc.ca
} ---------- } From: Timothy Moeller[SMTP:tmoeller-at-noran.com] } Sent: September 17, 1998 12:37 PM } To: Microscopy-at-Sparc5.Microscopy.Com } Subject: an apology (and a warning!) } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } At the risk of further crowding your in-basket(s), I want to apologize } to everyone for my recent mishap in posting an article (Re: Evaporation } of Gold) which resulted in bouncing ad infinitum (or ad nauseum, if you } prefer.) I believe the problem stems to a mistake I made in replying to } the list on that subject. Apparently, you are not supposed to use the } "Reply" function, but rather post a NEW message to the List Server at } {Microscopy-at-MSA.Microscopy.Com} . So consider this a warning in addition } to an apology -- it could happen to you as easily as it has happened to } me. In fact, I'd seen this same thing happen to other posters as well } recently, who apparently made the same mistake I did. } } ---------------------------------------------------------- } Timothy G. Moeller | Microanalysis Products } Senior Software Engineer | NORAN Instruments Inc., } {tmoeller-at-noran.com} | a ThermoSpectra company } ---------------------------------------------------------- }
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Hi, Luc et al.
Yes, having TWO (or more) tungsten filaments twisted together is important, to give the capillary action for more even dispersal as you mention. I forgot that important detail (it's been 27 years!) Thanks for jogging my memory. In fact, as I remember now, we were actually taking a long strand of tungsten wire and doubling it over more than once, to give a twisted set of 4 or more for our apparatus. We then wrapped the gold wire around that.
Regs, -- Tim
Luc Harmsen wrote: } Hi all. } Great topic. } We have been involved in building a new control unit for a factory which evaporates aluminium onto plastic parts to give them that silver look. } This has given us a lot more understanding of the coating technique as they have very large vacuum chambers and many samples that need an even and smooth coat each time. } What they do is to use tungsten wires coils that have two strands of W wire twisted together as aposed to, what we all seem to use, the single strand of W wire. The aluminium wire is then twisted by hand onto this coil, fairly loosely but just that it makes contact. The reasoning is that as you start the heating of the aluminium it will melt and flow, via capillary action, between the two W wires. This means that the aluminium is then in very good contact with the W coil and a lower current is needed to vaporise the aluminium. You also have a source of aluminium the whole length of the W coil each time. This ensures a even and repeatable coat each time. } } Cheers } Luc Harmsen } Anaspec, South Africa } International technical support on microscopy. } Tel: +27 (0) 11 476 3455 } Fax:+27 (0) 11 476 7290 } anaspec-at-icon.co.za } } -----Original Message----- } } From: Timothy Moeller [SMTP:tmoeller-at-noran.com] } Sent: Wednesday, September 16, 1998 7:15 PM } To: Paul.Gerroir-at-crt.xerox.com } Cc: Microscopy-at-sparc5.microscopy.com; Microscopy-at-sparc5.microscopy.com } Subject: Re: Evaporation of Gold } } Hi, Paul. } } Well, you've probably heard enough ideas on this subject. But I'll give } you my $0.02 worth (that's Au standard :-).) } } Working as a lab technician at UW Astrophysics Lab in Madison, WI, I } operated a vacuum deposition apparatus invented by a grad student which } vaporized gold wire and deposited it on the surface of insulated copper } wire to form the anode of a proportional detector. Anyway, in this } apparatus the gold wire was simply wrapped around tungsten (heater } filament) wire by hand. In vacuum, the tungsten filament was heated and } the gold wire melted, resulting in gold vapor being deposited evenly on } the wire rotissing nearby in the apparatus. It all seemed pretty crude } to me, but worked just fine. And I didn't take any special precautions } in wrapping the gold wire around the tungsten heater wire. } } Regs, } -- Tim } --- } } Gerroir, Paul J wrote: } } Hello Interested Readers, } } } } I have some experience in evaporating silver, (2 - 3mm shot) from a } } tungsten wire basket, however, I am now faced with challenge of evaporating } } Au wire onto a similar substrate. Is it appropriate to simply wrap the Au } } wire around the larger diameter tungsten wire and proceed or is there a } } better approach. Your comments/suggestions are appreciated. Thanks. } } } } Regards, } } Paul Gerroir } } Xerox Research Center of Canada }
-- .we now return control of your computer screen to you... ---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ---------------------------------------------------------- "I've spent my whole life trying to think up crazy ways of doing things." - Chief Engineer Montgomery "Scotty" Scott (TNG:"Relics") ----------------------------------------------------------
Hi: I am looking for a used Zeiss 10C or Zeiss 109 for a customer of mine. The scope will be used in the Los Angeles area. Please respond directly to me. Thank you. Peter Jordan, EMSI 909 694-1839
The pits and lands on a CD are { {inside} } the plastic of the CD, so you need to dissolve it with some sort of solvent (I believe I used methanol, but can't recall for sure). The correct organic solvent will make the plastic disappear entirely, not just craze it and turn it cloudy. I think there may be several types of plastic used as the base, because I used toluene a number of years ago the first time I tried this, and it didn't work on the CD I
used a few months ago. Anyway, use a couple of fresh changes until the foil is floating free, then mount it on a stub and take a look. You might need to mount both sides of a single piece so that you are sure you have the side with the pits. If you've gotten all the plastic off, you might get away without coating, but I coated mine anyway.
You can see an example of the pits and lands at http://www.mta.ca/~jehrman/cd.htm
BTW, the CD I used is a Microsoft(TM) Office demo CD, so any flaws are Bill Gate's fault, not mine!
Cheers,
Jim
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
Hello, all! This is proving to be a bit more "thorny" than I had expected. Thanks to those who have responded so far. Apparently there isn't any easy reference on TEM populations. Any or all additional inputs will be appreciated, and I WILL summarize and share the results! Thanks!
The repeating messages are NOT associated directly with an individual poster, so please don't complain to them or post your messages to the server it's not their fault as far as I can tell.
So far the only thing I have been able to determine for sure is that:
1.)" ALL " the repeating messages are definitely being bounced by a computer called "listserv.okstate.edu"
2.) "MOST" of those messages have "CC" microscopy
I have put the listserv.okstate.edu computer on the rejection list, which means if for some reason your Email routes through "okstate.edu" your posting may get rejected.
Do not "CC" microscopy. You should send all messages directly to our main address of
Microscopy-at-MSA.Microscopy.Com
while people have CC'ed Microscopy for several years, something has obviously changed recently (although not here) which causes CC'ed messaged which pass through listserv.okstate.edu to bounce.
My guess is that there is something strange going on at listserv.okstate.edu and I'm investigating. I seem to recall some recent requests from subscribers from that domain saying that their Email system has changed. Perhaps that is the source of the problem. I am investigating.
For now you will just have to bear with the problem.
The pits and lands on a CD are { {inside} } the plastic of the CD, so you need to dissolve it with some sort of solvent (I believe I used methanol, but can't recall for sure). The correct organic solvent will make the plastic disappear entirely, not just craze it and turn it cloudy. I think there may be several types of plastic used as the base, because I used toluene a number of years ago the first time I tried this, and it didn't work on the CD I
used a few months ago. Anyway, use a couple of fresh changes until the foil is floating free, then mount it on a stub and take a look. You might need to mount both sides of a single piece so that you are sure you have the side with the pits. If you've gotten all the plastic off, you might get away without coating, but I coated mine anyway.
You can see an example of the pits and lands at http://www.mta.ca/~jehrman/cd.htm
BTW, the CD I used is a Microsoft(TM) Office demo CD, so any flaws are Bill Gate's fault, not mine!
Cheers,
Jim
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
The pits and lands on a CD are { {inside} } the plastic of the CD, so you need to dissolve it with some sort of solvent (I believe I used methanol, but can't recall for sure). The correct organic solvent will make the plastic disappear entirely, not just craze it and turn it cloudy. I think there may be several types of plastic used as the base, because I used toluene a number of years ago the first time I tried this, and it didn't work on the CD I
used a few months ago. Anyway, use a couple of fresh changes until the foil is floating free, then mount it on a stub and take a look. You might need to mount both sides of a single piece so that you are sure you have the side with the pits. If you've gotten all the plastic off, you might get away without coating, but I coated mine anyway.
You can see an example of the pits and lands at http://www.mta.ca/~jehrman/cd.htm
BTW, the CD I used is a Microsoft(TM) Office demo CD, so any flaws are Bill Gate's fault, not mine!
Cheers,
Jim
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
If you've already checked the rollers in the Polaroid film back, the barring is probably on the CRT screen, and the next step is to calculate the frequency of the interfering signal. Simply divide the number of cycles of the barring in one scan line by the line time. If this comes out to 60 Hz, you probably have a ground loop, or an external magnetic field problem. If the frequency is 120 Hz, the problem is almost certainly ripple in the output(s) of your SEM's power supply. This may be a component failure (probably a capacitor, rectifier, or pass transistor), or could come from low line voltage. I've had problems with that on some older Philips machines, their power supplies really depend on line voltage being up to 100% of spec.
On Thu, 17 Sep 1998 micro-at-ldeo.columbia.edu-at-sparc5.microscopy.com wrote:
} Date: Thu, 17 Sep 98 13:33:55 EDT } From: "micro-at-ldeo.columbia.edu"-at-sparc5.microscopy.com } To: microscopy-at-sparc5.microscopy.com } Subject: sem image problems } } ------------------------------------------------------------------------ } 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 collegues, } } Does anyone know why some Polaroid shots from an SEM have faint parallel } vertical lines with a consistant spacing of somewhat less than 0.5 mm? Are } they in the film? I'm also getting patches of horizontal lines in some } micrographs lately that I'm pretty sure aren't due to either charging or } the Polaroid rollers. Could this be interference from the new lab next } door? } } Thanks, } } Dee } } } ____________________________________________________________________________ } Note: Sometimes I don't receive incoming emails (with no notification to } the sender). If I don't respond to your message, please send it again! } ____________________________________________________________________________ } _ } Dee Breger } Manager, SEM/EDX Facility } Lamont-Doherty Earth Observatory } Route 9W } Palisades NY 10964 USA } } T: 914/365-8640 } F: 914/365-8155 } I: www.ldeo.columbia.edu/micro } } }
Robert Wieland wieland-at-me.udel.edu Neither Yankee nor Dixie, east of the Mason-Dixon line (look it up). You can't go faster than light, you can't get colder than absolute zero, and you can't help somebody by not telling them the truth.
Dee Berger wrote: Does anyone know why some Polaroid shots from an SEM have faint parallel vertical lines with a consistant spacing of somewhat less than 0.5 mm? Are they in the film? I'm also getting patches of horizontal lines in some micrographs lately that I'm pretty sure aren't due to either charging or the Polaroid rollers. Could this be interference from the new lab next door?
Dee, in 1987 I observed parallel lines, about 0.3mm separation, in Polaroid Type 52 film. The lines ran at a slight angle to the horizontal. Because the only appeared in one lot number of film and not others I blamed the film. By this time, however, we did not have much of that lot left and chose to ignore it. I still have copies those photos. I have not observed them since, but we have not used Polaroid film in quite a few years. Try exposing the film on an optical microscope. If you see them again, send the film and some of the photos to Polaroid. If it is their film, I'm sure they would like to know. If you don't see the lines, they are probably being produced in the SEM.
Dennis B. Barr (dennbarr-at-eastman.com) Physical Chemistry Research Laboratory ECC Physical & Analytical Chemistry Research Division B-150B, R-132E, (423) 229-2188
} -----Original Message----- } From: "micro-at-ldeo.columbia.edu"-at-Sparc5.Microscopy.Com } [SMTP:"micro-at-ldeo.columbia.edu"-at-Sparc5.Microscopy.Com] } Sent: Thursday, September 17, 1998 1:34 PM } To: microscopy-at-Sparc5.Microscopy.Com } Subject: sem image problems } } ---------------------------------------------------------------------- } -- } 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 collegues, } } Does anyone know why some Polaroid shots from an SEM have faint } parallel } vertical lines with a consistant spacing of somewhat less than 0.5 mm? } Are } they in the film? I'm also getting patches of horizontal lines in some } micrographs lately that I'm pretty sure aren't due to either charging } or } the Polaroid rollers. Could this be interference from the new lab next } door? } } Thanks, } } Dee } } } ______________________________________________________________________ } ______ } Note: Sometimes I don't receive incoming emails (with no notification } to } the sender). If I don't respond to your message, please send it again! } ______________________________________________________________________ } ______ } _ } Dee Breger } Manager, SEM/EDX Facility } Lamont-Doherty Earth Observatory } Route 9W } Palisades NY 10964 USA } } T: 914/365-8640 } F: 914/365-8155 } I: www.ldeo.columbia.edu/micro }
I want to thank everyone who write to me about imaging a CD. The method that worked best (quickly, easily and no solvents!) was suggested by Brian McIntyre - see below. Just use a sticky tab on a stub to pull off the Al layer, sputter coat and scope. The images are nice. I also put some pieces of the CD in a solvent and got the Al layer that way, too. I apologize for ignoring the thread on CD prep recently. I do biological work and never thought I would need that info. So thanks to all who were willing to go there again.
best regards, Beth
} the surface you want to view is just under the top (label) side. the } easiest way to get the bit structure is to scribe a .5 X .5 cm square area } with a razor blade and then pull off the top plastic layer with the } underlying aluminum layer (with the bit structure replicated from the } bottom plastic layer) with a sticky tab on a metal stub.......this } technique works OK if you just want to see some pits and lands, it doesn't } however give a great whole piece sample. but this "problem" is surely } outweighed by its simplicity!! } } } good luck! } b- } } **************************************************************** } Brian McIntyre } Electron Microscopy Lab } Institute of Optics } University of Rochester } Rochester, NY 14627 } } 716-275-3058 } 716-244-4936(fax) } "Be well, do good work, and keep in touch"
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
Dear Timothy, } } Look, I'm sorry. But this wasn't my doing. True, it stems from an } oversight on my part (by posting to the wrong address, as I explained in } my apology article), but I did NOT compound the error myself by } re-sending the message, nor repeating the CC's, nor anything. The } listserver did all that. This whole thing just blew up in my face, } after I posted one simple little article incorrectly. I'm sorry, but } it's out of my control. } To err is human. To foul it up completely requires a computer. :-) Yours, Bill Tivol
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I have received a number of responses to my original message (enclosed below) desiring info on a digital camera for LM however most have been from vendors. I would really like information from users.
I want to amend my original message with an additional requirement for a digital camera. Since I want maximum versatility, I do not want a system that requires hook-up to a computer during capture. I want a camera that can be easily moved for use on different microscopes which may not be in the same room. This would mean I need an image storage system like a PCMCIA card which can be used to collect the images and then read at a computer at another location. Our computers are in a room across the lab from the LM rooms and I do not want to tie up a network capturing images and transfering from afar. We also certainly do not need additional computers, nor do I want to take up resources or space for a computer dedicated to this camera. We also are a MAC lab so can handle images in TIFF format best as this can easily be dealt with on either computer platform.
I imagine that these requirements will really limit our choices but am shooting for everything and will worry about compromising if necessary later.
Thanks for the responses, Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
Original message:
I am presently researching digital cameras which can be mounted on light microscopes for use for both fluorescence and bright field image capture. This is to augment film recording, not replace it. Resolution is very important as is even light spread. I am also interested in the different image storage mechanisms utilized by these cameras.
Any suggestions or personal experiences with using digital cameras for this purpose would be appreciated.
by nottingham.ac.uk with esmtp (Exim 1.92 #1) for Microscopy-at-Sparc5.microscopy.com id 0zJwpz-0000wY-00; Fri, 18 Sep 1998 10:28:23 +0100 Received: from CCW0F/SpoolDir by hermes.nottingham.ac.uk (Mercury 1.43); 18 Sep 98 10:28:24 GMT0BST Received: from SpoolDir by CCW0F (Mercury 1.43); 18 Sep 98 10:28:07 GMT0BST
Dear Microscopists I am looking for a bell jar (150mm diameter, 127mm high) for an Emscope SC500 coating unit. If any one has a spare they don't need and willing to part with please get in touch with me directly. We are able to pay for the item and shipping costs, but within the UK would be preferable.
Thanks a lot Nikki Nikki Bock EM Technician Dept. Materials Engineering University of Nottingham Nottingham NG7 2RD (0115) 9513759/9513871 Email: emznjb-at-hermes.nottingham.ac.uk
} We have several old American Optical/Spenser stereo microscopes that need a
} round glass stage insert. The inserts are 4" (10 cm) in diameter. I know
} we can have these made locally by a glass shop but was wondering whether
} anyone knows of an "off the shelf" source.
}
} Gary Radice
} Department of Biology
} Richmond VA
Gary,
These should probably be available through Leica. Also, you might want to contact a local service agent. Sometimes they know of sources which are more economical. If you don't have one in your area, contact me directly.
I recently had the same problem with faint parallel lines on Polaroid film with my SEM. This problem was not film-related and was simply due to electromagnetic interference from a nearby CRT. I had recently placed an X-terminal (with 20" monitor) on a table about 12 - 18 inches from the photo CRT on the SEM. The remedy was simple: either turn off the CRT when taking photos or move it away from the SEM console (3 - 4 feet was sufficient).
It sounds as though you may be having a similar problem and some kind of EM interference seems likely. Before shooting a Polaroid picture, try turning off devices around your SEM that are likely culprits, particularly any computer monitors, or ask the folks in the new lab next door if they will do the same. In lieu of that, you may be having an electronics problem with the microscope, such as high voltage instability, and your service person can help with that.
Hope this helps.
Dave
Dave Joswiak Dept. of Astronomy University of Washington Seattle, WA 98195
Hello collegues,
Does anyone know why some Polaroid shots from an SEM have faint parallel vertical lines with a consistant spacing of somewhat less than 0.5 mm? Are they in the film? I'm also getting patches of horizontal lines in some micrographs lately that I'm pretty sure aren't due to either charging or the Polaroid rollers. Could this be interference from the new lab next door?
Debbie Here's a summary of the most useful replies I received in response to my = inquiry about digital cameras. Naturally, most of the responses were from distributors, although many = had very informative content. Just the same, I left off their names in = the interest of fairness and maximizing S/N of this mailing list. I'm = happy to send unedited responses to anyone offline. Eric
------------ I think that there are two cooled CCD systems that you might consider. 1.) The first would be the Princeton Instruments MicroMax, a 12 bit, 5 = MHZ camera with a Grade 0 interline CCD, with thermoelectric cooling to = -10 C, which is also very fast as well, giving a full frame (1300 X = 1030) readout in 0.28 sec. This camera (including PCI interface) sells = for $19,500. 2.) An alternative system would be the Photometrics series 300, a 16 = bit, back illuminated camera using Grade 1, site 502B CCD, with = thermoelectric cooling to -25 C. This camera is not as fast as the = MicroMax, but has twice the QE, giving a full frame readout (512 X 512) = in 1. 4 sec, which may not be fast enough for your purposes. This camera = (including PCI interface) sells for $20,995. 3.) A third option would be the Quantix camera, also from Photometrics. = This is a 12-bit, 5 MHZ camera, using a Grade 1 KAF1400 CCD, also with = thermoelectric cooling (and the option for liquid cooling to -35 C). = This camera is similar in speed to the MicroMax, giving a full frame = readout (1317 X 1035) in 0.3 sec. This camera (with PCI interface) sells = for $24,495. One of the main differences between the first and third options is the = CCD chip. The interline chip is traditionally more sensitive in the = 'blue-green' region of the spectra than the KAF 1400 chip which is more = sensitive in the 'red' region of the spectra. Therefore if you will be = needing speed and sensitivity in the red as well as blue green part of = the spectra then the Quantix would be favored otherwise the MicroMax = would be favored. ---------- Olympus America sells a complete line of digital cameras for fura = applications. Currently we recommend either a frame transfer camera or = an interline camera for fura. These cameras are typically 12 - 14 bit = cameras with very good sensitivity for fura. Frame rates are variable, = typically from 1fps - 50fps. =20 You do not have to use ICCD cameras for fura. Digital cameras offer = several advantages over ICCDs. They have a greater dynamic range, = variable exposure rates and greater signal to noise ratio. ----------=20 I just received and quickly tested an Apogee KX2 - a cooled camera with = the Kodak 1300 chip. I am doing fluorescence work - although not = biological - and decided on this camera because of the dynamic range / = spectral range (I am using DAPI and IR filter sets) and price. Of = course it doesn't have quite the QE of a backilluminated camera, but on = first test it appears more than sufficient. Knowledgeable folks at that = company. Their first line of business is with astronomers but their = microscopy offerings are worth a look. Dave Calvert Eastman Chemical Co. P.O. box 1972 Lincoln Street=20 Kingsport, TN 37664 voice: (423) 229-4943 fax: (423) 229-4558 calvert-at-eastman.com ---------- I'm not sure what a "back-illuminated non-intensified" camera is, but I = would highly recommend a cooled CCD camera for your applications, = probably something like the Spot or maybe the Spot Jr., from Diagnostic = Instruments ($5-8K). =20 It depends on your application, but a 1/2 second exposure would be = pushing it for just about any CCD camera if you want color. There are = other high-end, high-speed digital cameras (maybe something from Dage = MTI) that could do this in B&W, but they are in the range of $20-25K.=20 ------------ We too are looking at digital cameras - had a demo of the olympus one = yesterday which I was impressed with! It did AO fluorescence on problem = - still have to try with other low light sources. problem with the olympus - nothing longer than 0.5s - and we regularly = take stuff of 2-3 s... ---------- Give our web on the new DVC 1300 digital cameras a review. It is very = detailed. http://members.aol.com/dvcco We can offer 1300 x 1030 pixels with integration / no cooling to 3 sec = or so with 10 bit S/N for a cost effective $4995 !!!! We are plug and = play with any RS-422 frame grabber board and we offer the boards also. = We are the US manufacturer of the camera and have taken on most board = lines for your convenience. ------------ Right now, the state of the art in Fura detectors is a Sony designed = chip family of progressive scan, interline transfer, Hyper HAD chips = like the ICX061 we use in our ORCA series cameras. These chips are used = by us and Princeton Instruments in cameras that produce high signal to = noise ratio images in low light conditions and are especially sensitive = in the blue wavelengths. The cameras cost about $15,000 and you will need a frame grabber and = software in addition. The total comes to about $20,000 usually. If = this is of interest to you, let me know and we will send you more = details. ------------ If you have to limit your exposures to 1/2 second, the SPOT camera will = not work as it is a real "Light Hog", and most exposures even with = bright fluorescence take 3-4 seconds per color channel. A color image = would then require 12-16 seconds to capture. Please call me at the = number below if you'd like to get pricing on some of our intensified CCD = cameras. We may be able to give you better pricing than you have been = seeing.=20 ----------- I sell a digital camera from the company Pixera. They did develop a = new digital camera. The sensitivity of this camera is 0.3 lux. You = can make frames accumulated or averaged. Max exposure time 12.8 sec. The complete system would cost about 6k. including a reduction lens for = the microscope.
For some work that we are doing we would like to know a source for Transine or Ammonium Fluoride. Does anyone know of a source? Thanks Al Bingham Semoptics Ltd. A manufacturer of Custom components for Scanning Electron microscopes.
I once saw a related problem in a Hitachi H2460N SEM, in which the Polaroid film would show lines at the top and bottom, but the middle would be clear. The lines faded out and back in again as the middle of the image was approached and passed. If memory serves me, the Hitachi service folks puzzled over this one and finally fixed it by adjusting the relationship of the X and Y screen scan controls. (If anyone from Hitachi is reading this maybe you can clarify/correct this?)
This particular artifact also showed up on the screen, however, while using the slower scan speeds.
Randy
Randy Tindall Electron Microscope Laboratory Box 3EML--Biology New Mexico State University Las Cruces, NM 88003
We are trying to embed relatively large (4mm x 5mm x 3mm) tissue samples in JB-4 and cut serial 5 micron sections for purposes of 3D reconstruction studies. The tissue of interest is the lamina cribrosa (connective tissue in eye). We are having a devil of a time for a variety of reasons, mostly related to my lack of expertise with JB-4. (For a complex set of reasons we cannot use paraffin.)
Can anyone help with the following questions? 1. The hardness of the resulting blocks is highly variable (maybe depending on the age of the catalyst?) The product insert states that more catalyst is needed as it ages, but gives no guidelines as to how much. Any ideas? 2. Sometimes the tissue sample becomes nearly transparent after infiltration, other times it does not. We cannot deduce why this occurs. 3. Some degree of curling and deformation of sections is inevitable. Any tips to reduce/minimize this?
Many thanks in advance, Ross Ethier -- Prof. C. Ross Ethier Department of Mechanical and Industrial Engineering, University of Toronto Toronto, Ontario M5S 3G8 email: ethier-at-mie.utoronto.ca voice: (416) 978-6728 fax: (416) 978-7753 http://mie.utoronto.ca/staff/profiles/ethier.html
I used to work in an ECM lab, so we had plenty of bottles of collagen lying about. Now that I'm ordering it for a new lab, I'm not sure if my predecessors just used what was available, or what was optimal.
When subbing slides (making them sticky so sections stay on), does it matter what sort of collagen is used? Is one particular type or bloom preferable,or just the cheapest? If the cheapest one is the answer, does anyone use Knox unflavored gelatin?
Thanks in advance.
Charlie Ginsburg NCC Research Dept. Lombard IL _________________________________________________________ DO YOU YAHOO!? Get your free -at-yahoo.com address at http://mail.yahoo.com
Dear Makroskopists, Someone asked me: what is the difference between the Wild Makroskop M420 and the M450? I know that the M420 is the M410 with with a trinoc or photo tube but I can find no info on the M450. Thank you.
########################### John Bozzola, Ph.D., Director Center for Electron Microscopy Southern Illinois University Carbondale, IL 62901 Phone: 618-453-3730 Fax: 618-453-2665 ###########################
I think the "Gold" bouncing Email is done. If your interested read on..
First the bouncing mail was NOT the fault of either the Microscopy Listserver or of Timothy Moeller {tmoeller-at-noran.com} . If any of you vented on Tim, you owe him an apology.
The whole problem was a computer system at OKSTATE.
What apparently happened is that the system administrator's at OK State decided to rename a Email node from okway.okstate.edu to osu-com.okstate.edu. They did this without notice of the users and without setting up forwarding of Email from old to new system ID's.
The computer okway.okstate.edu serviced the EM facility and the Microscopy Listserver had 3 subscribers there.
In their infinite wisdom the SysOp's somehow mis configured things and mail did not forward from okway.okstate.edu to osu-com.okstate.edu but rather created the loop. Basically, their POP3 Server, which kept trying to receive mail for the now defunct computer okway.okstate.edu kept appending Microscopy-at-... to the CC: list of the mail which could not be delivered to the 3 people at the EM facility. Then, rather than just dying like a broken Email pipe sh ould have done, their's server forwarded mail to the CC list as if it originated at okway. This sent the mail back to Microscopy via the CC, which inturn sent it back to okway etc, etc, etc,.... hence the loop.
I would like to find out if and where there is an agent for PZO, the Polish Optical Company (other than in Poland). I am not having any luck corresponding directly and wish to contact an agent.
I think possibly the M450 was of the same series as the popular M400 but included some sort of built in illuminator. I am not positive but can probably look it up some time this week. If you have not already obtained an answer I can probably let you know sometime this week. -----Original Message----- } From: John J. Bozzola {bozzola-at-siu.edu} To: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com}
The only digital camera for LM that does not require to be permanently connected to a computer I have heard of, is Olympus DP10. It uses SmartMedia card, although a permanent computer connection is an option. We do not have the camera, but considering to purchase it.
Alex ______________________________ Alexander Titkov
Millennium Inorganic Chemicals PO Box 245 Bunbury WA 6231 Australia Ph: (08) 9780 8505 FAX: (08) 9780 8500 E-mail: atitkov-at-micl.com.au
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I have received a number of responses to my original message (enclosed below) desiring info on a digital camera for LM however most have been from vendors. I would really like information from users.
I want to amend my original message with an additional requirement for a digital camera. Since I want maximum versatility, I do not want a system that requires hook-up to a computer during capture. I want a camera that can be easily moved for use on different microscopes which may not be in the same room. This would mean I need an image storage system like a PCMCIA card which can be used to collect the images and then read at a computer at another location. Our computers are in a room across the lab from the LM rooms and I do not want to tie up a network capturing images and transfering from afar. We also certainly do not need additional computers, nor do I want to take up resources or space for a computer dedicated to this camera. We also are a MAC lab so can handle images in TIFF format best as this can easily be dealt with on either computer platform.
I imagine that these requirements will really limit our choices but am shooting for everything and will worry about compromising if necessary later.
Thanks for the responses, Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
Original message:
I am presently researching digital cameras which can be mounted on light microscopes for use for both fluorescence and bright field image capture. This is to augment film recording, not replace it. Resolution is very important as is even light spread. I am also interested in the different image storage mechanisms utilized by these cameras.
Any suggestions or personal experiences with using digital cameras for this purpose would be appreciated.
I'd really have to see the micrographs, but the vertical variations sound like some sort of electronic problem, although the frequency seems abnormally high. Can you provide more information in regards to the total record time and the number of lines of resolution used? Possibly the filtering of the accelerating voltage (usually around 20KHz) or the PMT high voltage (around the same frequency). In either case, the frequency would produce a vertical banding that would not be strictly vertical but at least slightly diagonal.
The horizontal variations may be a problem with the stage electrical connection. The sample is grounded through a connection that must remain constant, but in older instruments may vary. Normally a copper or copper-berylium spring contact is used that rubs against the sample holder through sample rotation. In older instruments, that contact may be flakey due to buildup of contaminants or reduction of spring force. Find the connection and clean both the spring loaded contact and the rotational surface it contacts with and try to reform the spring to produce a stronger contact.
} Hello collegues, } } Does anyone know why some Polaroid shots from an SEM have faint } parallel vertical lines with a consistant spacing of somewhat less } than 0.5 mm? Are they in the film? I'm also getting patches of } horizontal lines in some micrographs lately that I'm pretty sure } aren't due to either charging or the Polaroid rollers. Could this be } interference from the new lab next door? } } Thanks, } } Dee } } } ____________________________________________________________________ } ________ Note: Sometimes I don't receive incoming emails (with no } notification to the sender). If I don't respond to your message, } please send it again! } ____________________________________________________________________ } ________ _ Dee Breger Manager, SEM/EDX Facility Lamont-Doherty Earth } Observatory Route 9W Palisades NY 10964 USA } } T: 914/365-8640 } F: 914/365-8155 } I: www.ldeo.columbia.edu/micro } } } Allen R. Sampson Advanced Research Systems 317 North 4th. Street St. Charles, IL 60174 PH 630.513.7093 FAX 630.513.7092 Email: ars-at-mcs.net WWW: http://www.mcs.net/~ars Analytical instrument maintenance services
Ken, I think you're wrong here. Silicone based oils will crack into an electrically non-conductive form in an electron optics column. The resultant contaminant coating will cause sample and optics component charging which will affect the electron beam. In my work, I suggest and use only Santovac DP oils, not just for their cracking characteristics to an electrically conductive film but also for their tolerance of large influxes of air while heated.
The ETEC instruments switched over, when properly calibrated, to the diffusion pump at 70 microns. This is appropriate to the range of a diffusion pump. But many other instruments switch over earlier, sometimes as high as 150 microns. In those SEMs, the early switch-over results in a stalling of the diffusion pump and a large release into the chamber of diffusion pump oils. As a case in point, I offer the Hitachi S-570. The preferred cure is a delay in the diffusion pump switch over to well under 100 microns.
Even in a well calibrated SEM, the delays in switching over to the diffusion pump and moderate air leaks can lead to a stalled diffusion pump. While sudies have shown that the primary contribution to most contamination problems is from mechanical pump oils, my practical experience is that contamination is often from diffusion pump oils. This also naturally leads to the question of what is an appropriate oil to use for mechanical pumps. Another thread?
} steve-at-facstaff.wisc.edu wrote: } } Dear all } } } } I may be dragging the thread away from contamination, but I } } thought that most electron microscope users avoided silicon based } } oils in diff pumps etc } } } } because they are extremely difficult to remove from the interior } } of an electron microscope and any contamination would normally } } have electrical insulating properties (catastrophic in an e.m.). } } Perhaps I am wrong but I would welcome any comments. After all } } this is one of the reasons why Santovac oils and their relatives } } became so popular (despite their costs). } } } } If I am labouring under a mis-apprehension then I apologise. } } } } Malcolm Haswell } } University of Sunderland } } UK } } } } All of the silicon based DP fluids I am aware of (which may not be } } all that are on the market) will break down under an electron beam } } and deposit a layer similar to glass on the nearest cool surface } } in the column. I don't know of any EM manufacturers that would } } recommend their use because they are almost impossible to remove } } if they do get in the column. We don't even use silicon based } } fluids in our vacuum evaporator. } } } } K. A. Brackett, Ph.D. } } TN & Assoc./USEPA } } } } Steve Limbach } } Associate Researcher } } Bock Research Lab. } } 1525 Linden Dr. } } } } UW-Madison } } Madison, Wisc. 53706 } } } } TEL 608 263-2582 } } FAX 608 262-4570 } } EMAIL slimbach-at- facstaff.wisc.edu } } Malcom, } I've been watching this thread with considerable interest, also. } ETEC sold almost every one of its microscopes with Transene Vacoil } (later Vacoil-S) which I believe is redistilled Dow-Corning 705. I } have never had any problems with it in 21 years of servicing these } instruments. Even a burped DP (and I've dealt with my share) has } never been a problem beyond the fact that you have to clean } everything. Many ETECs were sent out with the water-cooled baffle } plumbed in just before the DP rather than before the power supplies. } Those instruments will condense MP oil on their EDS detectors until } they get replumbed. but silicone doesn't seem to present any } problems. } } Ken Converse } owner } Quality Images } third party SEM service } Delta, PA } Allen R. Sampson Advanced Research Systems 317 North 4th. Street St. Charles, IL 60174 PH 630.513.7093 FAX 630.513.7092 Email: ars-at-mcs.net WWW: http://www.mcs.net/~ars Analytical instrument maintenance services
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} Dee, I had a recurring problem of vertical lines on the SEM image and Polaroids that was caused by noise in the digital logic circuits. This SEM had these logic circuits installed but not used. They would occasionally generate the vertical lines and the fix was to clean the circuit board "fingers" with and eraser. The boards were those in the CRT imaging bin. Hope this helps. Dave Audette audette-at-osi.sylvania.com } }
PZO has a website, with a possibility to send them an email trough that chanel. I have had some "bounced messages" some time ago. Perhaps the problem is solved now.
http://www.PZO.com
You'll find a link to their website in english there.
I don't know about PZO dealers in your country, their agent in Germany (and probably Europe...) is:
Gerhard G=F6ke Bahnhofstr. 27 D-58095 Hagen Germany. Telefon 02331/31754 Telefax 02331/31754
Dee, I have found the old Polaroids which exhibited parallel lines similar to those you wrote about. THEY WERE OPTICAL MICROGRAPHS. So the lines couldn't have been due to electronic interference. They were spaced at ~0.3 micron and were found throughout the image. We tried several different Polaroid film holders and got the same results with all of them, so we concluded the problem was not due to the film rollers. When we switched lots of film the problem went away.
Has anyone else ever seen lines like this?
Dennis B. Barr (dennbarr-at-eastman.com) Physical Chemistry Research Laboratory Physical & Analytical Chemistry Research Division Eastman Chemical Company Kingsport, TN 37662-5150
Debby - you are looking for a digital camera to suit your particular needs. It's a good starting point, but it is equally important to see what is available. Manufacturers of these instruments balance carefully customer's needs with what is technically possible and desirable. Professional digital cameras generally and those for microscopy in particular are tethered for good reasons. The non-tethered cameras for micrography most likely are toys. The problem is not just image storage, but extensive pre-photography software functions that are not available in any conventional camera. Most importantly the computer's screen serves as a quite superior viewer in all of these systems and, somehow, that requires a computer close to the camera. I cannot foresee this changing in the medium term. You could have a "roving" digital camera for your microscopes, by using a laptop, perhaps with a "real" screen attached. I expect, however, that in the long term it would be more practical to move the microscopes and to utilise a centrally located computer between a couple of superior microscopes. For class use, several 1 or 2 mega-pixel cameras could give students full-on experience in the new skills required with digital photography. The other points made: Uneven illumination relates to lenses or condenser system used and has nothing to do with the digital camera. Only "toy cameras" do not have the option to produce files in TIFF. Our online catalogue has a lot of non-camera specific digital photography information, particularly on the /epix.htm page I was not one of the vendors who wrote to Debby offering a digital camera, but would like to make the point that most vendors have much experience and most are honest. Sure, they may present a bias viewpoint, but don't expect to get an unbiased view from users either. They made a decision to buy a particular brand and they have experience with a particular range of jobs using that particular brand, well or badly. You can only hope for a diversity of views and that you have good judgement to pick A suitable system. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au *****
I have received a number of responses to my original message (enclosed below) desiring info on a digital camera for LM however most have been from vendors. I would really like information from users.
I want to amend my original message with an additional requirement for a digital camera. Since I want maximum versatility, I do not want a system that requires hook-up to a computer during capture. I want a camera that can be easily moved for use on different microscopes which may not be in the same room. This would mean I need an image storage system like a PCMCIA card which can be used to collect the images and then read at a computer at another location. Our computers are in a room across the lab from the LM rooms and I do not want to tie up a network capturing images and transfering from afar. We also certainly do not need additional computers, nor do I want to take up resources or space for a computer dedicated to this camera. We also are a MAC lab so can handle images in TIFF format best as this can easily be dealt with on either computer platform.
I imagine that these requirements will really limit our choices but am shooting for everything and will worry about compromising if necessary later.
Thanks for the responses, Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
Original message:
I am presently researching digital cameras which can be mounted on light microscopes for use for both fluorescence and bright field image capture. This is to augment film recording, not replace it. Resolution is very important as is even light spread. I am also interested in the different image storage mechanisms utilized by these cameras.
Any suggestions or personal experiences with using digital cameras for this purpose would be appreciated.
Dee, On my SEM if I use the quick scan to produce an image I get lines. When I ues the slow scan the lines are gone. It has to do with the way the recording monitor displays the image. Have you tried the different photo rates on the scope?
-- Regards, Gregory Rudomen Technical Specialist University Microscopy Imaging Center State University of New York at Stony Brook 516-444-3126 Greg-at-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. **********************************************************
Well, I guess that system is not OK after all. yuk, yuk, yuk.
Thanks Nestor for taking care of it!
Paula :-)
} Colleagues... } } I think the "Gold" bouncing Email is done. If your interested read on.. } } First the bouncing mail was NOT the fault of either the Microscopy } Listserver or of Timothy Moeller {tmoeller-at-noran.com} . If any } of you vented on Tim, you owe him an apology. } } The whole problem was a computer system at OKSTATE. } } What apparently happened is that the system administrator's at } OK State decided to rename a Email node from okway.okstate.edu to } osu-com.okstate.edu. They did this without notice of the users } and without setting up forwarding of Email from old to new system } ID's. } } The computer okway.okstate.edu serviced the EM facility and the } Microscopy Listserver had 3 subscribers there. } } In their infinite wisdom the SysOp's somehow mis configured } things and mail did not forward from okway.okstate.edu to } osu-com.okstate.edu } but rather created the loop. Basically, their POP3 Server, which } kept trying to receive mail for the now defunct computer okway.okstate.edu } kept appending Microscopy-at-... to the CC: list of the mail which could not be } delivered to the 3 people at the EM facility. Then, rather than just } dying like a broken } Email pipe sh ould have done, their's server forwarded mail to the CC list } as if } it originated at okway. This sent the mail back to Microscopy via the CC, which } inturn sent it back to okway etc, etc, etc,.... hence the loop. } } Once again we have been done in by someone else. } } Cheers.... } } Nestor } Your Friendly Neighborhood SysOp }
Paula Sicurello UC Berkeley Electron Microscope Lab psic-at-uclink4.berkeley.edu phone: 510-642-2085 fax: 510-643-6207 http://biology.berkeley.edu/EML
It was a great pleasure hearing Nancy Lane, a Cambridge cell biologist, speaking of the wonders of the microscope (including the electron variey) on the Classic FM programme "Masters of Their Art" (Saturday 19th September). This is Britain's largest commercial radio station.
Her work in particular uses invertebrates to research systems which have parallels in ourselves, such as the cell membranes that constitute the blood-brain barrier. Invertebrates generally reach us, at best, on the table, and so the "cruelty and cuddle" factors do not as yet restrict their use in experiments. But she deplored the low profile invertebrate studies have in biology courses, likening it to the situation where people talk of "shellfish" in a restaurant without regard to their nearly inexhaustible variety.
Still, such ignorance would be easily dispelled by a trip to Hong Kong, where I for one have expanded my gastronomic experience by at least two PHYLA, the echinoderms (sea cucumber) and the coelenterates (fried jellyfish).
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
At 09:00 AM 9/21/98 -0400, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I'm looking back a few years here, but I seem to recall a problem that was once caused by the rubber rollers in the Polaroid film holder (the one for P/N 55), rather than the metal rollers which squeeze the chemical packet. These rubber rollers had finely spaced parallel ridges (don't know the width) that can harden with encrusted chemicals if the unit is not frequently cleaned.
This doesn't answer as to why switching film lots caused the problem to go away, unless perhaps one lot had the quality of being ultra-sensitive to pressure. Anyone who has ever dropped a Polaroid film on the floor and stepped on it has seen the tread pattern of their sneakers on the final image.
This is probably reaching way out for an explanation, but your message did ring a bell. For what it's worth.
Randy
Randy Tindall Electron Microscope Laboratory Box 3EML--Biology New Mexico State University Las Cruces, NM 88003
Hi Charlie, we have always used the following mixture to coat slides so that plant sections (from paraffin embedments) stick:
Haup'ts adhesive
one gm Knox gelatin dissolved in 100ml dH2O at 30C. After dissolution of the gelatin add 2 gm phenol and 15 ml glcerin. Filter.
The phenol is added to prevent fungus from colonizing the gelatin but, as phenol is not nice, I have lately dispensed with it. Instead, freeze the mixture in aliquoits in Eppendorf tubes that can be thawed as required. Coat slides by rubbing 1-2 drops of the mixture onto the slide and letting sit until dry. Do not do this with bare fingers if there is phenol in the mix or ever if cleanliness is a concern. Hope this helps, John
} } I used to work in an ECM lab, so we had plenty of bottles of collagen } lying about. Now that I'm ordering } it for a new lab, I'm not sure if my predecessors just used what was } available, or what was optimal. } } When subbing slides (making them sticky so sections stay on), does it } matter what sort of collagen is used? } Is one particular type or bloom preferable,or just the cheapest? If } the cheapest one is the answer, does anyone use Knox unflavored gelatin? } } Thanks in advance. } } } Charlie Ginsburg } NCC Research Dept. } Lombard IL
________________________ C. John Runions Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
I have not seen MgO now for eight years, but I used to see a lot of them, together with other refractories. We compared grain-size by light microscopy. The problem with polishing with diamond paste only is that you get a perfect flat surface, without clear indications of grain boundaries, at least if you are dealing with 96+ % MgO, not the less pure ones. This prefect flat surface is probably what you mean by "smearing", because you expected grain boundaries. A simple method to make part of the grain boundaries visible is to do a last polish NOT with diamond paste, but something softer, such as alumina or aluminosilicate, the softer the better (and around 1 micron or finer). This gives relief polishing, boundaries becoming visible by differences in orientation . However, not all grain boundaries will be well visible, some boundaries have to be "filled in" by the observer.
A better visiblity of grain boundaries is provided by a thermal etch, but even here there is not 100% visibility of the grain boundaries. Saw a 1-2 mm section with the polished surface from the MgO (remove embedding material) and put that for 1/2 - 1 hour in a furnace at about 700C. These conditions are what I remember after 8 years, so I may be off slightly. With best regards, Emond de Roever
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I have been asked to compare the grain size of 2 MgO aggregates. The samples were embedded and polished with diamond paste, but I am not happy with the result. The surface looks smeared. Does the sample need to be etched or do I need to evaluate my polishing technique???? Thanks for you help.
Nan Laudenslager Specialty Minerals, Inc. Easton, PA nhl-at-early.com
by is2.nyu.edu (8.8.8/8.8.7) with SMTP id PAA01370; Mon, 21 Sep 1998 15:45:41 -0400 (EDT)
On Mon, 21 Sep 1998, Robert H. Olley wrote:
} Her work in particular uses invertebrates to research systems which have } parallels in ourselves, such as the cell membranes that constitute the } blood-brain barrier. Invertebrates generally reach us, at best, on the } table, and so the "cruelty and cuddle" factors do not as yet restrict } their use in experiments. But she deplored the low profile invertebrate } studies have in biology courses, likening it to the situation where people } talk of "shellfish" in a restaurant without regard to their nearly } inexhaustible variety.
I don't know your experience but invertebrates provide the substrate for much of classic and present research in basic neurobiology. And the advantages they provide are greatly appreciated and extolled in the standard texts.
} Still, such ignorance would be easily dispelled by a trip to Hong Kong, } where I for one have expanded my gastronomic experience by at least two } PHYLA, the echinoderms (sea cucumber) and the coelenterates (fried } jellyfish).
Or to modern Chinese and Japanese restaurants in the States.
} } I used to work in an ECM lab, so we had plenty of bottles of collagen } } lying about. Now that I'm ordering } } it for a new lab, I'm not sure if my predecessors just used what was } } available, or what was optimal. } } } } When subbing slides (making them sticky so sections stay on), does it } } matter what sort of collagen is used? } } Is one particular type or bloom preferable,or just the cheapest? If } } the cheapest one is the answer, does anyone use Knox unflavored gelatin?
If you simply want to make your slides sticky for paraffin or plastic sections, here is a protocol that works a lot better than polylysine, chromgel, or anything else I have ever used. It is also cheap!
PREPARATION OF COATED SLIDES Protocol 05.01.02 - last updated 9/5/95
Reference: In situ hybridization to cellular mRNAs using radioactively labeled RNA probes. L. Angerer (1989) In: In situ hybridization: methods for detecting DNA and RNA sequences at cellular and subcellular resolution. Am. Soc. Cell Biol. Workshop Manual. pp 1-21. Based on the more complex technique of Gotlieb and Glaser (1976) BBRC 63:815-821.
HAZARDOUS CHEMICALS: Do not use this protocol if you are not aware of all the safety precautions required to work with these chemicals. Consult all material safety data sheets. Work in a fume hood with proper safety technique. Wear appropriate safety gear.
SUPPLIES: 3 Coplin jars; 1 stock bottle; forceps. They need to be clean and lint free. Wear gloves when making the solution and dipping the slides.
1. Make a 2% 3-aminopropyltriethoxysilane(Sigma) working solution. Put 98 mls of acetone in the Schott bottle (orange cap) that is dedicated and marked for this purpose. Add 2 ml of 3-aminopropyltriethoxysilane. Pipet up and down a few times to mix the solution.
2. Fill the first Coplin jar with enough stock solution to cover the slide. Put the lid on unused stock. As the 3-aminopropyltriethoxysilane evaporates, refill Coplin jar with fresh stock. Fill a second Coplin jar with approximately 50 mls of acetone. Fill a third Coplin jar with approximately 50 mls of dH2O.
3. Line the work space in the fume hood with paper towels and place about three or four test tube racks on top of them as rests to lean the slides against after dipping.
4. To begin dipping the slides, you will need a pair of forceps. First, place between five and seven slides in the first jar (the acetone/aminopropyltriethoxysilane working solution). Wait about one minute before removing them.
5. Using the forceps, transfer these slides to the second Coplin jar (acetone). While you are letting these slides sit in the acetone for a minute, you can begin putting new slides into the first jar. If one is preparing large batches of slides, replace the acetone rinse periodically.
6. Transfer the slides in the second jar to the third jar (dH2O). Again, you will want to let the slides sit in the jar for about a minute. While waiting for these slides, you can continue to add slides to the first and second jars. If one is preparing large batches of slides, replace the dH20 periodically.
7. Before removing the slides out of the dH2O, you will want to dip them up and down a few times to make sure they have been rinsed well. Using the forceps, lean the slides against the test tube racks in order for them to dry.
8. Repeat this process until you have coated sufficient slides. When the slides are dry (about 10-15 min.), store them in a labeled slide box. The slides should be clear. White spots are a sign of improper rinsing.
9. When you are finished dipping the slides, clean up the hood. Do not save the stock solution longer than a couple of hours. Place the stock solution from the stock bottle and first Coplin jar into the appropriate hazardous waste bottle. Rinse these two containers with the acetone from the second Coplin jar. Discard the acetone into the appropriate hazardous waste bottle. You may dispose of the dH2O in the sink. The three Coplin jars and stock jar should be placed neatly off to the side within the hood to await future use. 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 (573)-882-4712 (voice) (573)-882-0123 (fax) --============_-1305711192==_ma============ Content-Type: text/enriched; charset="us-ascii"
} } I used to work in an ECM lab, so we had plenty of bottles of collagen
} } lying about. Now that I'm ordering
} } it for a new lab, I'm not sure if my predecessors just used what was
} } available, or what was optimal.
} }
} } When subbing slides (making them sticky so sections stay on), does it
} } matter what sort of collagen is used?
} } Is one particular type or bloom preferable,or just the cheapest? If
} } the cheapest one is the answer, does anyone use Knox unflavored gelatin?
If you simply want to make your slides sticky for paraffin or plastic sections, here is a protocol that works a lot better than polylysine, chromgel, or anything else I have ever used. It is also cheap!
{bold} {bigger} {bigger} PREPARATION OF COATED SLIDES
{/bigger} {/bigger} {/bold} Protocol 05.01.02 - last updated 9/5/95
Reference: In situ hybridization to cellular mRNAs using radioactively labeled RNA probes. L. Angerer (1989) In: In situ hybridization: methods for detecting DNA and RNA sequences at cellular and subcellular resolution. Am. Soc. Cell Biol. Workshop Manual. pp 1-21. Based on the more complex technique of Gotlieb and Glaser (1976) BBRC 63:815-821.
HAZARDOUS CHEMICALS: Do not use this protocol if you are not aware of all the safety precautions required to work with these chemicals. Consult all material safety data sheets. Work in a fume hood with proper safety technique. Wear appropriate safety gear.
SUPPLIES: 3 Coplin jars; 1 stock bottle; forceps. They need to be clean and lint free. Wear gloves when making the solution and dipping the slides.
1. Make a 2% 3-aminopropyltriethoxysilane(Sigma) working solution. Put 98 mls of acetone in the Schott bottle (orange cap) that is dedicated and marked for this purpose. Add 2 ml of 3-aminopropyltriethoxysilane. Pipet up and down a few times to mix the solution.
2. Fill the first Coplin jar with enough stock solution to cover the slide. Put the lid on unused stock. As the 3-aminopropyltriethoxysilane evaporates, refill Coplin jar with fresh stock. Fill a second Coplin jar with approximately 50 mls of acetone. Fill a third Coplin jar with approximately 50 mls of dH2O.
3. Line the work space in the fume hood with paper towels and place about three or four test tube racks on top of them as rests to lean the slides against after dipping.
4. To begin dipping the slides, you will need a pair of forceps. First, place between five and seven slides in the first jar (the acetone/aminopropyltriethoxysilane working solution). Wait about one minute before removing them.
5. Using the forceps, transfer these slides to the second Coplin jar (acetone). While you are letting these slides sit in the acetone for a minute, you can begin putting new slides into the first jar. If one is preparing large batches of slides, replace the acetone rinse periodically.
6. Transfer the slides in the second jar to the third jar (dH2O). Again, you will want to let the slides sit in the jar for about a minute. While waiting for these slides, you can continue to add slides to the first and second jars. If one is preparing large batches of slides, replace the dH20 periodically.
7. Before removing the slides out of the dH2O, you will want to dip them up and down a few times to make sure they have been rinsed well. Using the forceps, lean the slides against the test tube racks in order for them to dry.
8. Repeat this process until you have coated sufficient slides. When the slides are dry (about 10-15 min.), store them in a labeled slide box. The slides should be clear. White spots are a sign of improper rinsing.
9. When you are finished dipping the slides, clean up the hood. Do not save the stock solution longer than a couple of hours. Place the stock solution from the stock bottle and first Coplin jar into the appropriate hazardous waste bottle. Rinse these two containers with the acetone from the second Coplin jar. Discard the acetone into the appropriate hazardous waste bottle. You may dispose of the dH2O in the sink. The three Coplin jars and stock jar should be placed neatly off to the side within the hood to await future use.
i've been doing some analysis of particulate matter filtered from urban and rural air using the old PRC program that tracor-northern sold in the '80s (boy do i feel old!). i have a few definition files setup to classify the particles, but i'm wondering if anyone out there has some files that work well for them that they'd like to share; i'd like to try some "better" or at least different classification schemes to see what i get. (this work is being done as part of a class project in environmental engineering for freshman undergraduates....its a hoot!)
thx!
b-
**************************************************************** Brian McIntyre Electron Microscopy Lab Institute of Optics University of Rochester Rochester, NY 14627
716-275-3058 716-244-4936(fax) "Be well, do good work, and keep in touch"
Hi, Recently I was quite unable to email the listserver. Some users may have had the same experience, so it seems worth posting. My attempted postings were returned with various computer generated messages, most commonly: "554 MX list for sparc5.microscopy.com points back to ultra.ultra.net.au" or "local configuration error"
"Points back", to me meant that the listserver, for whatever reason refused the email. I contacted Nestor via postmaster-at-microscopy.com with "I am not a spam" in the subject line. The short of it is that our service provider had changed their software about 10 days earlier. Amazingly, only about four out of dozens of email addresses caused problems but the fault clearly lay with our service providers software. Happily our genial and patient (going by emails only!) SysOp pointed me in this right direction. Thank you Nestor. Cheers Jim Darley ProSciTech Microscopy PLUS PO Box 111, Thuringowa QLD 4817 Australia Phone +61 7 4774 0370 Fax: +61 7 4789 2313 Great microscopy catalogue, 500 Links, MSDS, User Notes **************************** www.proscitech.com.au *****
} In theory, I expect that when compared with a tungsten } filament, at least the solid, single post LaB6 cathode } design by Kimball, would make for less movement, especially } during the warming-up period.
Just for the sake of fairness among us vendors, let me point out that both Denka (the LaB6 manufacturer which we represent in the United States) and FEI make LaB6 cathodes in which the crystal is mounted on rigid molybdenum posts, and this design is also recommended for microanalysis by virtue of its mechanical stability.
{snip}
Jim Darley added: } The obvious way to increase stability in LaB6 is to } increase the size of the microflat at the tip of the LaB6 } cone. The normal size of the cone (Kimball's) is 15 or 20 } micrometer. A 40 micrometer microflat is also available } (unfortunately at greater cost) and this makes a LaB6 quite } suitable for quantitative microanalyses. Brightness is } reduced but for microanalyses that is no consideration.
Again, this is product-specific. Denka, for example, produces "microflats" of 20, 40, 60 and 100 microns. It is also important to take cone angle into consideration. Both 60 and 90 degree cone angles are available, and the 60 degree cone angle has proven very useful for applications, like electron beam lithography, which require maximum stability.
Best regards, Steven E. Slap
******************************** Energy Beam Sciences, Inc. The Laboratory Microwave Company http://www.ebsciences.com ********************************
Jim Darley wrote} In theory, I expect that when compared with a tungsten } filament, at least the solid, single post LaB6 cathode } design by Kimball, would make for less movement, especially } during the warming-up period. } In practise, tungsten filaments are more stable because the } emitting area is much larger and so, minor misalignment due } to a drifting filament is less consequential. } The stability we are talking about is due to slow drift and } this is of no consequence to normal, including high } resolution imaging in TEM or SEM. Stability over several } minutes matters when performing quantitative microanalyses } with a probe or EDS in dedicated SEM/TEM. In quantitative } analyses a spectrum maybe acquired over two minutes and } that is related to standard spectra and all numbers must } lead to results that come within 1% of reality. } The obvious way to increase stability in LaB6 is to } increase the size of the microflat at the tip of the LaB6 } cone. The normal size of the cone (Kimball's) is 15 or 20 } micrometer. A 40 micrometer microflat is also available } (unfortunately at greater cost) and this makes a LaB6 quite } suitable for quantitative microanalyses. Brightness is } reduced but for microanalyses that is no consideration. } Reducing downtime by increasing "filament life" to over } 5000 hours at low and medium emission is the main benefit } of a Lab6 in a microprobe. } For non-analyses work, a standard flat Lab6 cathode is } quite stable for SEM and TEM requirements; extreme drift } would matter but stability worries in these applications } are more likely to relate to HV instabilities or a pulsing } beam due to column/aperture contaminations. The question } for Mark Darus is the EM's vacuum system: Is it good enough } for LaB6 operation. } Disclaimer: ProSciTech supplies Kimball cathodes and } filaments.
******************************** Energy Beam Sciences, Inc. The Laboratory Microwave Company http://www.ebsciences.com ********************************
We use Fisherbrand Superfrost/Plus Microscope Slides from FisherScientific. Cat # 12-550-15. They are more expensive than plain glass ones, but you save the trouble of having to mix up the collagen or p-L-lysine, worry about storage, or deal with contamination by fungus, etc. Sections never fall off, yet if you want to do in situ embedding, you can still get the resin slab off. I highly recommend them.
No commercial interest; just a satisfied customer.
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
I'll add my two cents regarding the discussion of Uranyl acetate radiation. Two years ago our Radiation Safety Office in Beltsville, MD asked our EM group to be responsible for the storage, etc. of all uranium compounds located at our site. I had heard the same comments in the past about not "needing to worry about UAc radiation". "Hey, the old reagent bottles don't even have radiation warnings on them"! "There is only a slight amount of alpha radiation".
Anyway, while gathering all uranium compounds, our radiation safety officer checked the compounds using a dosimeter that recorded alpha, beta, and gamma radiation. Levels of radiation were found that were just below OSHA guidelines for maximum daily exposure when the powder was checked from several inches away from the dosimeter. Thus, we decided to store the compounds in an acrylic box behind a beta shield in an isolated location. Whenever the actual Uranyl acetate is weighed to prepare dilute solutions proper safety precautions are recommended, i.e. use a beta shield, wear gloves and dust mask, weigh in a low occupancy room, etc.
We repeated our dosimeter readings last month with two different alpha, beta, and gamma dosimeters taking readings several feet from the bottles, removing the two acrylic shields one by one, and then with the cover of the UAc exposed (always moving closer to the chemical source). I won't enter the millirem/rad discussion; suffice to say it's an interesting little experiment for your support staff especially with the dosimeters left on audio signal. The dilute solutions themselves don't generate much radiation above background levels but should be treated as heavy metal wastes (don't pour down sinks).
Thanks for now.
PS Vote for most dangerous EM chemical DAB (diaminobenzidene).
Bruce F. Ingber Biologist- Electron Microscopy USDA-ARS, SRRC 1100 Robert E. Lee Blvd. New Orleans, LA 70124
} ---------- } From: William Tivol[SMTP:tivol-at-wadsworth.org] } Sent: Wednesday, September 16, 1998 10:48 AM } To: michowej-at-nus.edu.sg } Cc: microscopy-at-Sparc5.Microscopy.Com } Subject: Re: Uranyl Acetate radiation } } ------------------------------------------------------------------------ } 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 Josephine, } } } } A solution of 10g UA in 15mls H2O was measured with a Geiger counter. } 500 } } counts/sec was generated. } } A supplier had measured 100g UA :- } } 1 Alpha - {2 counts/sec, using a 540 scintillation meter with AP-2 Probe } } 2 Beta - } 500 counts/sec, using a 540 E1 probe coupled to a GM Meter (this } } determines beta events and some low energy gamma events) } } 3 Gamma dose Rate (energy field) - two measurements done: } } using Mini monitor tpye R with GM Probe - 0.6mR/hr (mainly gamma) } } and Ionisation chamber DMM 95/0500 - 5 mR/hr (Beta and Gamma energy } } field). } } 4 Specific Activity (U approx. 55%) = 1.04 x 10 { {...} } Bq { {...} } gm } } { {...} } . } } } I calculated approximately the expected activity from 30 g UA } (about 0.1 mole, or 6*10^22 atoms). T_1/2 is 4.4*10^9 y and there are } 3.1*10^7 s/y, so the decay rate is 5*10^-18 s^-1, and the activity is } 3*10^5 Bq. } The build-up of daughter products with shorter half-lives will } reach steady state at which point the activities of the daughters will be } the same as that of the parent. Pa 234 has a gamma transition, and there } are several betas in the chain. The longer-lived isotopes in the chain } have lives of 10^4 to 10^5 y, and these will not be at steady state (unless} } your UA is *very* old ;-) ). 0.6 mR/hr is a significant amount of exposure, } and, if one were to hold the jars for some minutes, a sizable fraction of } the allowed annual dose would be attained. } } } Can UA be used openly without protection in laboratory? } } } Small amounts can be used, but be sure to wash hands before eating. } One area of the lab should be used for UA. A quiet area with little traffic } is best. UA, while not nearly the most dangerous EM reagent, should still } be treated with respect. } Yours, } Bill Tivol }
i have a collegue that just got a hold of a relatively old ( about 20 yrs ) spectrophotometric detector from perkin elmer model LC-75, however the opreating manuals seem to be missing. I realize that this is not a chromatography listserve, however, i was wondering if someone had any information that could help him out. He's contacted the manufacturer and they only sell manuals. He's a little hesitant to start purchasing material for this unit until he is certain that it can be of use to him.
thanks in advance
Michael Mandanas Particulate Materials Center 218 MRL Bldg. Pennsylvania State University University PArk, PA 16802 mxm67-at-email.psu.edu
Untethered, no cables required, camera, with high resolution? Full color?
There are VERY few. The best ones seem to be collaborations by Kodak with Nikon and Canon ( http://www.kodak.com/global/en/professional/products/cameras/range.shtml and http://www.kodak.com/US/en/digital/genInfo/EOSDCS1DCS460.shtml ). The Kodak DCS 460 (NIKON N90 camera body) and the Kodak EOS-DCS 1 (Canon EOS-1N body) were designed (1) to match the resolution of 35mm ISO 80-100 film; (2) be usable by professional photographers as to get the benefits of digital photography with the functionality and feel of an SLR camera. The Recording CCD records at 2036 x 3060 x 36bit (i.e. 18MB tiff files) with a sensitivity of ISO 80 (which is a little low for fluorescent work but usable). Both cameras utilize regular Nikon or Canon lenes, as well as the respective camera mounts for microscopes. "The images you shoot are stored on small, removable storage cards. A 170 MB card holds up to twenty-six 18 MB images. When you fill up one card, simply load another and keep shooting.After downloading image files from a storage card to your computer, you can even shoot over old files. Record more than 200 images per battery charge or an unlimited number if you're using the AC battery charger/adapter. " (You can get PC or Mac adapters to plug directly into your computer system.) Or you can capture directly via SCSI cable.
Sounds like what we're all looking for in a general LM camera? The problem is price Last time I got a price quote they were ~$13k. There are others available from Kodak with lower resolutions as well as high senesitivity (i.e ISO 200 - 1600 ), and I seemed to recall having read that Kodak is working on a new camera with both the high 2k x 3k resoltuion and high light senesitivity.
Sorry, since I still can't afford one, I can't tell you how well they handle LM. (Oh, I don't work for any vendor, I'm not selling anything, nor do I own stock in Kodak - at least I don't think I do.... : ).
So you see Virginia there is a Santa Claus (He's just really expensive).
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
"640K ought to be enough for anybody." -- Bill Gates, 1981
Botanical Microtechnique and Cytochemistry by G.P. Berlyn and J.P. Miksche. Iowa State U. Press. 1976. ISBN 9138-0220-2
Nice book! Note, this is my copy, I don't know if there is a more recent edition, but Books In Print (at your local library or bookstore) should have the current information (you may need to check Forthcoming Books In Print). I used this text among others when I learned microscopy.
Phil
} Can anyone give me some references that pertain specifically to botanical } microtechniques? } } Preferably in english... } } Thanks! } } Yvan Lindekens.
}}}}}}}}}}}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{{{{{{{{{{ Philip Oshel PO Box 620068 Middleton, WI 53562 (608) 833-2885 oshel-at-terracom.net or poshel-at-hotmail.com
Subbing slides. Make huge quantities at once using nothing but gelatin and water. (Additives may interfere with the stability of LM stains which engage in redox shifts with alacrity). Dry the slide, store dustfree, and throw out the gelatin mixture. Do not keep gelatin mixture even overnight. No problem with bacteria ever this way. The slides last forever. (0.1% gelatin in water is good, more is better ifyou can tolerate it. The thicker the gelatin coat the more interferece with photomigrography, but the more secure the section on the slide). So long, Hildy
i've been doing some analysis of particulate matter filtered from urban and rural air using the old PRC program that tracor-northern sold in the '80s (boy do i feel old!). i have a few definition files setup to classify the particles, but i'm wondering if anyone out there has some files that work well for them that they'd like to share; i'd like to try some "better" or at least different classification schemes to see what i get. (this work is being done as part of a class project in environmental engineering for freshman undergraduates....its a hoot!)
thx!
b-
**************************************************************** Brian McIntyre Electron Microscopy Lab Institute of Optics University of Rochester Rochester, NY 14627
716-275-3058 716-244-4936(fax) "Be well, do good work, and keep in touch"
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We recently had an interesting problem with vertical even spaced lines. We added a digital aquisition system to our JEOL JSM-840 SEM. All worked fine until we tried to capture an image using the compositional backscattering detector. The evenly spaced vertical lines appeared across the entire image. They did not appear under TOPO-BSI or SEI imaging of the same area.
Further investigation showed that the lines were still there, although the spacing changed, even when filament heating and KV were turned off as long as the mode selector was in COMP. It turned out that the problem was a wire from the COMP-SEI detector to ground. This wire had been there since the purchase of the microscope and we never could see a problem when recording COMP-SEI images using film. It only showed it's ugly side when recording using digital acqusition. The wire was removed and all is well.....not sure why it was there in the first place since there was other grounding as well. I do have a wonderful example of electrical interference for my SEM class so all is not a loss.
Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
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Hi,
We are planning to use DAB and nickel sulfate methods to colocalize synaptic proteins in wet sections. Question: If two proteins occur in the same area, does one stain cover the other? Has anyone had this experience? Does anyone have a better idea using wet sections and a light microscope (rather than a confocal, etc)? I would so much appreciate any opinions. Bye, Hildy {hcrowley-at-du.edu}
Greetings fellow microscopists; I have recently inherited a Leitz Dialux 22EB microscope, with attached camera. While I have little trouble with basic operation of a light microscope, the camera and setup are unfamiliar to me... is there anyone out there with the same or similar model who still has the manual, and would be willing to send me a copy? I have tried contacting Leica, who seem to have either been Leitz in a past life, or absorbed them somehow, but have not received a response so far. Any help would be most appreciated.
thanks in advance shea
Dr. S. Shea Miller Agriculture & Agri-Food Canada Eastern Cereal & Oilseed Research Centre Rm 2068, Bldg 20, CEF Ottawa, Ontario Canada K1A 0C6 Phone: (613)759-1760 Fax: (613)759-1701 e-mail: millers-at-em.agr.ca
Hello fellow microscopists; I have inherited a Leitz Dialux 22EB light microscope, with an attached camera that is unfamiliar to me. The camera body says Wild MPS12 (Heerbrugg, Switzerland). While I suspect that I could eventually figure out how most of the stuff on the camera works (having been forced to do it with other scopes in the past) it would save me all kinds of time if someone has a manual kicking around that they could copy the pertinent bits for me from.
thanks in advance shea Dr. S. Shea Miller Agriculture & Agri-Food Canada Eastern Cereal & Oilseed Research Centre Rm 2068, Bldg 20, CEF Ottawa, Ontario Canada K1A 0C6 Phone: (613)759-1760 Fax: (613)759-1701 e-mail: millers-at-em.agr.ca
The books that I use are john e. sass Botanical Microtechnique Plant Microtechnique by Johansen I will look for extra johansen... thought I had an extra one here...
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many months ago a collegue of mine asked to the microscopist from this listserver about tips and receipes por treating sand grains covered with a biofilm builded up by bacteria.
The procedure we followed was designed according to the literature and some of the answers we got from some of you. At the same time we followed instructions from someone who had done something like that before here. Neither of both gave us good results. My question is what we did wrong?????
These are some details of the 2 procedures we followed:
Procedure I
1) Primary fixation with glutaraldehyde 2,5% in buffer phosphate pH=6,7, at refrigerator temperature, 3 hs.
2) Three washes 10 or 15 minutes each in buffer phosphate
3) Secondary fixation with OsO4 2% in H2O, in the same buffer, at the same temperature, 1 h in darkness.
4) Dehydration in ETOH 50%, 70% and 95% once, 10' each, and 100%, twice or three times for 15' each.
5) Drying by filtering under vacuum on a 45 microns sterile Micropore filter. (we don't have facilities for crytical point drying)
Procedure II
The same steps except for the 3), that means only one fixation step (as done by someone else here before)
What we got is something strange (at least to us, who don't have any experience on treating biological samples). The grains treated according to Procedure II look very clear, transparent in some cases, and only a few are dark or grey. We think that is normal since the fixation treatment is not the appropiate one. Those treated by Procedure I look from light grey to dark grey and even black, just like the grains collected from the bioreactor that treats wastewater from a dairy products manufacture.
Nevertheless, on the SEM the biofilm supossed to be added on the grain surface is gone on both samples!!!! Where has it gone? Why that difference in colour then?
Please, we need to hear comments about this because we have no idea what has happened.
One more detail, the glutaraldehyde 25% we use looks light yellow, its pH is 3,7. If that is the problem, too much polimerysation, why do we still get dark grains?
Thanks to all of you in advance.
Silvia Montoro Centro Regional de Investigacion y Desarrollo de Santa Fe Santa Fe Guemes 3450 3000 Santa Fe Argentina csedax-at-arcride.edu.ar fax +54 42 550944
The wire was removed and all is well.....not sure why it was there in the first place since there was other grounding as well. I do have a wonderful example of electrical interference for my SEM class so all is not a loss.
Debby Sherman, Manager Phone: 765-494-6666 Microscopy Center in Agriculture FAX: 765-494-5896 Dept. of Botany & Plant Pathology E-mail: sherman-at-btny.purdue.edu Purdue University 1057 Whistler Building West Lafayette, IN 47907-1057
This may be a silly question. What is the smallest object ( using brightfield illumination) that can be observed. For example, I calculated resolution of .4 micron for a 100x, .90 NA objective and wavelengh value of 540. This is the smallest separation between features that can be resolved. Is this also the smallest feature that can be observed given a single feature on a smooth background?
Michael Ingram Rodel, Inc Polishing Lab 451 Bellevue Rd Newark, DE 19713 (302) 366-0500, ext.: 2545
Seems to me that I have seen in one of the Popular Science type magazines that a company called Imagek owned by Irvine Sensors Corp. is going to sell something called an EFS-1. It is a drop in cartridge for your 35mm camera body that is battery powered and capable of capturing mega pixel images. It will come with an adapter to upload your images to a computer and I think they were planning to sell for under $1,000. The neat thing is that you could still use all those expensive camera optics that you still have around. As usual I have no financial interest in this product, just something I thought I'd interject into this thread. I just checked and they have a web site at http://www.imagek.com/index.shtml.
-- ================================================================== Greg Strout Electron Microscopist, University of Oklahoma WWW Virtual Library for Microscopy: http://www.ou.edu/research/electron/www-vl/ e-mail: gstrout-at-ou.edu Opinions expressed herein are mine and not necessarily those of the University of Oklahoma ==================================================================
Smallest object depends on contrast: much smaller (than .4 micron) hole in thin specimen with transmitted light illumination will be visible.
Vladimir Dusevich
On Wed, 23 Sep 1998, Ingram, Mike 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 may be a silly question. What is the smallest object ( using } brightfield illumination) that can be observed. For example, I } calculated resolution of .4 micron for a 100x, .90 NA objective and } wavelengh value of 540. This is the smallest separation between } features that can be resolved. Is this also the smallest feature that } can be observed given a single feature on a smooth background? } } Michael Ingram } Rodel, Inc } Polishing Lab } 451 Bellevue Rd } Newark, DE 19713 } (302) 366-0500, ext.: 2545 } }
This may be a silly question. What is the smallest object ( using brightfield illumination) that can be observed. For example, I calculated resolution of .4 micron for a 100x, .90 NA objective and wavelengh value of 540. This is the smallest separation between features that can be resolved. Is this also the smallest feature that can be observed given a single feature on a smooth background?
Michael Ingram Rodel, Inc Polishing Lab 451 Bellevue Rd Newark, DE 19713 (302) 366-0500, ext.: 2545
This is not a silly question, and I have seen a suprising number of people get confused by this. Here is a simple test: go outside at night and look at the stars. Now, individual stars are below the resolution limit of the unaided eye (ex. you won't be able to resolve a binary star system). However, obviously you can see (detect!) them. This is because resolution limits and detection limits are two different things. Think of a single small bright object on a dark background. As the object approaches a true point, the image approaches the point spread function (by definition). There is no reason to expect that as the object drops below some (resolution) limit, the light coming from the object will stop propagating to the detector. As long as it is there is enough light reaching the detector, you can still detect it. The image doesn't get smaller (since the PSF is the limit, and it will get dimmer, but it still can be detected). Resolution simply involves seeing how close _another_ object can get to the first one without their images overlapping by some amount (that amount depending on whether you use the Rayleigh or Sparrow criterea). The perception or detection of a bright object on a dark background is limited by the "brightness" of the object. Now, the perception of a dark object on a bright background is different situation. Consider a dark line on a bright background: what happens when the line narrows and the two line spread functions get close to start to overlap? The minimum will become shallower and shallower. Based on the eyes' ability to discern intensity variations, Pluta (reference below) gave formulas for the detection of these objects: Using typical parameters (488nm, 0.9NA) the smallest dark objects on a bright background that can be observed are 41nm for a disk, and 2.6nm for a line.
Resolution, perception/detection and location are different, but unfortunately all three tend to get lumped together as "resolution". I know you didn't ask, but since it's related, you can also locate an isolated small object to better than the resolution limit (if you know that your system has at worst only symmetric aberrations). Try this: draw a circle and then try to find the center. Remember, this is not a random processes, but a deterministic one. This also extends to edge location. One study (can't find the reference right now) back in 1986 showed that confocal microscopes (of that era) had a 20nm uncertainty in locating edges (the same value as for the SEMs of the day). This means that as long as a the two sides of a line object are resolved, then you can measure the width of the object to much better than the resolution limit. Obviously SEMs can perform the measurements on narrower objects than LM, but I did included the qualification concerning the object being wide enough for the edges to be resolved in the previous sentence. (No flames for saying that LM will replace EM: I stated the limitations twice! (and pointed it out again)) Again, resolution, perception/detection and location are different (related to the imaging system, but still different).
Regards, Matt Atkinson 3M Corporate Research Labs
(Pluta's book Advanced Light Microscopy, vol 1, pg337-348 gives a very good explanation.)
mingram-at-rodel.com wrote:
} This may be a silly question. What is the smallest object ( using } brightfield illumination) that can be observed. For example, I } calculated resolution of .4 micron for a 100x, .90 NA objective and } wavelengh value of 540. This is the smallest separation between } features that can be resolved. Is this also the smallest feature that } can be observed given a single feature on a smooth background? } } Michael Ingram } Rodel, Inc } Polishing Lab } 451 Bellevue Rd } Newark, DE 19713 } (302) 366-0500, ext.: 2545
Can anyone point me in the right direction concerning the preparation of inorganic colloid samples in moderate/high ionic strength waters for quantitative analysis by TEM? The dispersions contain relatively low number concentrations of particles which we are trying to measure.
So far, samples have been prepared by ultracentrifuging onto carbon films (based on published method of Nomizu et al. Electron microscopy of nanometer particles in freshwater, Anal. Chem. vol 60, 2653-2656, 1988) and wicking the residual solution away with a filter paper. The problem is that there are considerable drying artefacts such as matting of particles, uneven dispersion of particles within a grid square (possibly due to surface tension effects from residual water films during drying?) and crystallisation of residual salts.
I'd be most interested to hear from anyone with experience of preparation of this type of sample or any good ideas.
Thanks in anticipation,
Simon
Dr Simon Dumbill Team Leader, Materials Characterisation AEA Technology 220, Harwell Tel: +44 1235 434245 Didcot Fax: +44 1235 435941 Oxfordshire OX11 0RA Email: Simon.Dumbill-at-aeat.co.uk UK
May I humbly suggest the following recipe: 0.5% gelatin (300bloom, nothing fancy), 0.05% chrome alum. Clean slides with a good detergent, rinse extensively with tap, then d.i.H2O. Sprinkle gelatin on cold H2O, warm carefully until gelatin is dissolved (do not boil!), add chrome alum*. Do NOT add the chrome alum before the gelatin. You may dip the slides in warm or cool. Allow to air dry, loosely covered from dust. I do not store the solution.
I do use the fancy Fisher slides for in situs, but NOT for free floating sections. Free floaters are thicker than what is recommended for those slides and sometimes do not adhere well to them. *chrome alum=chromium potassium sulfate
Thank you Caroline for the message. Coincidentally, I have two stereo SEM books for children 8 years and up coming into the book stores in November. One is called "Bug Eyes and Butterfly Wings" and the other is called "Snail Tongues and Spider Fangs."
They come with stereo glasses. Big kids should enjoy them too! My co-authors are Shar Levine and Leslie Johnstone who have published quite a number of Science books for kids.
ISBN: 1-894042-16-6 and 1-894042-05-0 Publishers: Canada - Somerville House, USA - Andrews McMeel, New Zealand - David Bateman
If anyone is interested in SEM puzzles, check out "Small Wonders" on -at- Discovery Canada on the Discovery Channel every Monday night 8pm-9pm. Unfortunately, I don't think -at- Discovery Canada is shown anywhere but Canada, but you can still participate in the puzzle by going to the Discovery Channel web site at www.exn.ca. Then go to -at- Discovery and then "Small Wonders." It is an interesting web site for all sorts other science stories.
} } Are any of you interested in the possibility of using your micrographs to } illustrate children's books? Here's a conference that should be quite } informative: } } Marine Biological Laboratory Hosts Institute for } Children's Book Authors and Illustrators } October 9-11, 1998 } } Woods Hole, MA-The Marine Biological Laboratory's Science Writing } Fellowships Program and the Center for Children's Environmental } Literature is co-sponsoring an Author, Illustrator, Biologist } Institute. Working and aspiring children's book authors and } illustrators, as well as scientists, are invited to participate in the } three-day meeting. Organizers hope to foster new collaborations between } authors, illustrators, and biologists. } } } For more information, contact: } Pamela Clapp Hinkle } Director of Communications } Marine Biological Laboratory } 7 MBL Street } Woods Hole, MA 02543 } Tel: 508-289-7276 } Fax: 508-457-1924 } e-mail: pclapp-at-mbl.edu } } For a complete program, see http://www.mbl.edu/html/MISC/AIB.html. } } Caroline Schooley } Educational Outreach 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/Books.html } Intertidal invertebrates: http://www.fortbragg.k12.ca.us/AG/PCI/pci.html
Dr. Elaine Humphrey Biosciences Electron Microscopy Facility University of British Columbia 6270 University Blvd, mail-stop Botany Vancouver, BC CANADA, V6T 1Z4 Phone: 604-822-3354 FAX: 604-822-6089 e-mail: ech-at-unixg.ubc.ca
Some years ago I heard an account of a radiation safety inspection as part of a larger safety review at a large lab. The lab was involved in coal research, as were we, and had a fair amount of coal samples stored in glass jars. An inspection team came thru and happened to check the jars for radiation, found some, and instructed the researchers that they would need to start following radiation safety procedures.
Now coal can contain minute amounts of uranium in its mineral matter, but not enough that should set off a detector. Besides the detectors were setup to meaure alpha particles, and there was no way that alpha particles emitted from the contents of a glass jar should be detected on the outside.
A little digging revealed that there was in fact a little radiation present, but it was a slight residue left on the jar surface after washing. Apparently the jars went through the same washer as did other jars which had held radioactive materials. I think the radiation safety folks may have received additional training after the incident.
} Anyway, while gathering all uranium compounds, our radiation safety officer checked the compounds using a dosimeter that recorded alpha, beta, and gamma radiation. Levels of radiation were found that were just below OSHA guidelines for maximum daily exposure when the powder was checked from several inches away from the dosimeter. Thus, we decided to store the compounds in an acrylic box behind a beta shield in an isolated location. Whenever the actual Uranyl acetate is weighed to prepare dilute solutions proper safety precautions are recommended, i.e. use a beta shield, wear gloves and dust mask, weigh in a low occupancy room, etc. } } We repeated our dosimeter readings last month with two different alpha, beta, and gamma dosimeters taking readings several feet from the bottles, removing the two acrylic shields one by one, and then with the cover of the UAc exposed (always moving closer to the chemical source). I won't enter the millirem/rad discussion; suffice to say it's an interesting little experiment for your support staff especially with the dosimeters left on audio signal.
{snip}
---------------------------------------------------- Warren E. Straszheim 23 Town Engineering Iowa State University Ames IA, 50011-3232
Osmium is a contrast agent hence your black grains. As to the disapperance of the biofilm, I suspect that it is still there (at least to some degree), but that you cannot recognize anything biological because everything has completely collapsed/debrided during the final dehydration (essentially air-drying made even worse by the filtration under vacuum). I doubt you will see anything meaningful unless you find a critical-point dryer, particularly if the biofilm is rather thick to begin with. Bacterial cells are rather hardy, but there are good reasons why critical-point drying and other fancy methods were developed.
On the other hand, just what exactly are you trying to show by doing the SEM? If it is simply the presence of a biofilm, then why not use epifluorescence microscopy using some vital stains? It is a lot simpler and will give you more meaningful images than will SEM.
Rob Palmer CEB/UT
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Ingram, Mike wrote: } } This may be a silly question. What is the smallest object ( using } brightfield illumination) that can be observed. For example, I } calculated resolution of .4 micron for a 100x, .90 NA objective and } wavelengh value of 540. This is the smallest separation between } features that can be resolved. Is this also the smallest feature that } can be observed given a single feature on a smooth background? } } Michael Ingram } Rodel, Inc } Polishing Lab } 451 Bellevue Rd } Newark, DE 19713 } (302) 366-0500, ext.: 2545
-- Not a silly question; I have a similar question along these lines... Correct me if I'm wrong, but don't we usually just use the Rayleigh criterion as a rule of thumb for estimating resolution limit in any optical system such as this?
---------------------------------------------------------- Timothy G. Moeller | Microanalysis Products Senior Software Engineer | NORAN Instruments Inc., {tmoeller-at-noran.com} | a ThermoSpectra company ----------------------------------------------------------
} Can anyone point me in the right direction concerning the preparation } of inorganic colloid samples in moderate/high ionic strength waters for } quantitative analysis by TEM? The dispersions contain relatively low } number concentrations of particles which we are trying to measure. } } So far, samples have been prepared by ultracentrifuging onto carbon } films (based on published method of Nomizu et al. Electron } microscopy of nanometer particles in freshwater, Anal. Chem. vol } 60, 2653-2656, 1988) and wicking the residual solution away with a } filter paper. The problem is that there are considerable drying } artefacts such as matting of particles, uneven dispersion of } particles within a grid square (possibly due to surface tension } effects from residual water films during drying?) and } crystallisation of residual salts.
I work on similar things and have a number of methods.
Firstly, you can just proceed by dipping a copper grid with supported carbon film into the colloid (if fairly dilute in particles) or a diluted version of the colloid. The carbon film is then left to dry on some filter paper. This still gives problems of clumping and crystallisation of salts on drying but may be the only way in some cases.
Alternatively, one can filter the particles from the colloid (if they will filter and not just go straight through the filter paper), wash with water, and redisperse in an organic solvent such as methanol. Then dip a copper grid with a supported carbon film into the suspension. This avoids the salt problem and does give better particle dispersion on the grid although it is not perfect and some particle clumping always occurs. The concentration of the suspension is critical here: too high and lots of clumping is inevitable, too low and the concentration of particles on your carbon film is rather low. Only trial and error will give the best concentration for your taste.
Finally, one could centrifuge the particles out, pour off the aqueous solution, add methanol or ethanol and redisperse and proceed as above. This is an alternative for the case where the particles are impossible to filter off.
Hope this helps.
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ Ian MacLaren, Tel: (44) (0) 121 414 3447 IRC in Materials for FAX: (44) (0) 121 414 3441 High Performance Applications, email: I.MacLaren-at-bham.ac.uk The University of Birmingham, or: ianmaclaren-at-hotmail.com Birmingham B15 2TT, http://web.bham.ac.uk/I.MacLaren England. ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
After your final 100 ETOH rinse, try putting the samples in HMDS (hexamethyldisilazane) for about 30 minutes then keep in a desiccator for another 24 hours before viewing.
kim
} } 4) Dehydration in ETOH 50%, 70% and 95% once, 10' each, and 100%, twice or } } three times for 15' each. } } } } 5) Drying by filtering under vacuum on a 45 microns sterile Micropore filter. } } (we don't have facilities for crytical point drying) } }
} } } } Silvia Montoro } } Centro Regional de Investigacion y Desarrollo de Santa Fe } } Santa Fe } } Guemes 3450 } } 3000 Santa Fe } } Argentina } } csedax-at-arcride.edu.ar } } fax +54 42 550944 } }
HILDEGARD CROWLEY wrote: } } Hi, } } We are planning to use DAB and nickel sulfate methods to colocalize } synaptic proteins in wet sections. Question: If two proteins occur in } the same area, does one stain cover the other? Has anyone had this } experience? Does anyone have a better idea using wet sections and a light microscope (rather than a confocal, etc)? I would so much appreciate any opinions. } Bye, } Hildy } {hcrowley-at-du.edu}
Hildy:
I hate to say it all depends but it all depends. I have had good luck with doing DAB+Ni-Co first then DAB 'plain' for the second Ab but I was staining two different cell populations. When I was doing 2 Ab on one cell my results were mixed. Poor results when doing 2 monoclonals in one cell, good results when doing one mono and one poly. In an area as small as a synapse you might want to use two fluorescent markers, this usually gives very good results and can be very beautiful.
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 e-mail: mcauliff-at-umdnj.edu ***************************************************************
Anybody have a good protocol for preparing coccolithophores for taking pix of the intact coccospheres? I have fumbled around and obtained a few pix, but the yield of coccospheres is miniscule; these organisms seem to want to shuck their liths if you look at them cross-eyed.
I have a polarizer which is delaminating from the base glass substrate. This is in an rotating assembly specifically built as an Analyser for an older optical microscope.
Does anyone know how to either repair this or know where replacements can be obtained? I am assuming that the polarizer material is a linear polarizer - is this a fair assumption?
FYI: The polarizing material appears to be a thin platic-type material which has been glued to an underlying glass substrate. This sandwich is then mounted between 2 outer glass plates which have been (AR?) coated (or are these 1/4 wave plates?).
Any great ideas on how to recover this? Much thanks in advance - Ian L.
Wow! Thanks to everyone who responded so speedily to my cry for help. Sorry about the dual message... I tried to recall the first one, as it was only a day or so since I had tried to reach Leica via their web site. Leica (in the person of Philip Hyam) was in fact quite speedy in contacting me and assuring me of the availability of the manual.
Hi everybody! For one of our projects we used what we called crustacean buffer: 500mM NaCl, 12 mM KCl, 12mM CaCl2, 20 Mm MgCl2, 10 mM Tris (no HCl), and 4.3 mM Maleic Acid. We made double strenght buffer and mixed it with 4% aqueous sol. of osmium to obtain 2% working solution We did the same with glutaraldehyde: double strenght buffer with 10% aqueous glut to get 5% glut. Osmium solution truned yellow to brown within 1 hour after mixing and glut change color to yellow overnight. Needless to say fixation of the tissue was not great. Is there an expanation for that fenomena? I found a reference of maleate buffer used in staining procedures but not for fixation. Thank you. Dorota
} This may be a silly question. What is the smallest object ( using
} brightfield illumination) that can be observed. For example, I
} calculated resolution of .4 micron for a 100x, .90 NA objective and
} wavelengh value of 540. This is the smallest separation between
} features that can be resolved. Is this also the smallest feature that
} can be observed given a single feature on a smooth background?
}
} Michael Ingram
} Rodel, Inc
} Polishing Lab
} 451 Bellevue Rd
} Newark, DE 19713
} (302) 366-0500, ext.: 2545
Dear Michael,
The typical formula for resolution,R = (1.22 x lambda)/(NA objective + NA condenser), takes into account the following information:
(1) the nature of the background illumination, (2) orientation of the feature, (3) spacing (in your case, edge to edge) and (4) edge fidelity. Your calculation is correct for the smallest object which can be RESOLVED using brightfield microscopy and the optics quoted. However, providing that the object could scatter light (a pit, for instance) much smaller objects could be OBSERVED, i. e., you could tell that they were there but nothing about size, shape, etc.
"Optimizing Light Microscopy..." has a detailed but easily understood discussion of the whys and wherefors. More info is available on our website: { {http://www.MME-Microscopy.com/education} . (Don't be scared away by the fact that the book was originally written for biologists and clinicians. There's lots of pertinent stuff for materials scientists, too.).
In July I asked this group for assistance in getting Leica to respond to us. I would like to thank all of you that responded and to let you know that I heard from them soon after my message was posted. I would also like to thank Ed Rae for resolving the problem. I also apologize to Leica for taking this long to let you all know that they helped us out.
Sincerely Marty Reed Equipment Technician Biology Department Humboldt State University Arcata CA 95521 707-826-3234 707-826-3201 FAX mmr7001-at-axe.humboldt.edu
We would like to have regular object in the background to track cell movement relative to. We have used polystyrene beads in the past, but these are very difficult to work with and expensive. Someone in materials science recommended Lycopodium powder which I gather is some kind of dried fungal spore. Does anyone know how large and/or regular these spores are? Any alternative suggestions? We are looking for 0.5-5um and (obviously) easily visualized. Thanks- Dave Knecht
Dr. David Knecht Department of Molecular and Cell Biology University of Connecticut 75 N. Eagleville Rd. U-125 Storrs, CT 06269 Knecht-at-uconnvm.uconn.edu 860-486-2200 860-486-4331 (fax)
For the first clue, look at the objectives. If they are all 160 mm (a marking on the objective barrel like 160/0.17), they can be interchanged. If there is a sign for "infinity" (a figure 8, on its side) instead of 160 on both, they can be interchanged. The only problem arises when one is "fixed tube length" (160mm) and the other is "infinity corrected".
Re: Confocal - better check with Olympus. The hardware fitting and the optical planes may or may not be compatible.
Dear Simon, } } Can anyone point me in the right direction concerning the preparation } of inorganic colloid samples in moderate/high ionic strength waters for } quantitative analysis by TEM? The dispersions contain relatively low } number concentrations of particles which we are trying to measure. } What gives the dispersion its high ionic strength, and do you want to analyse it? If you are only interested in the particles, and if the high IS is not from volatile compounds, you will have to rinse the particles in H2O (or some volatile buffer). If you want to analyse the liquid phase, you will have to get areas where it is on the grid, but no particles.
} So far, samples have been prepared by ultracentrifuging onto carbon } films (based on published method of Nomizu et al. Electron } microscopy of nanometer particles in freshwater, Anal. Chem. vol } 60, 2653-2656, 1988) and wicking the residual solution away with a } filter paper.
This seems to be a good start. If you do not wick away all of the solution, you could plunge-freeze the grid and analyse the frozen-hydrated specimen, then freeze-dry in situ and re-analyse. Do the first on Friday in a cryo-holder (which must be kept full somehow), then raise the temp to ~-90 C. Leave under vacuum for 24 hrs, raise temp to -80 C, leave for another 24 hrs, raise to -70 C, leave for 24 more hrs, then do the analysis.
} The problem is that there are considerable drying } artefacts such as matting of particles, uneven dispersion of } particles within a grid square (possibly due to surface tension } effects from residual water films during drying?) and } crystallisation of residual salts. } The cryo procedure should take care of all but the crystallization problem (unless the residual salts are volatile--then they will be gone too).
} I'd be most interested to hear from anyone with experience of } preparation of this type of sample or any good ideas. } No experience with this procedure, but it was featured in an EDS class I took -at- Lehigh. Possibly some of the Lehigh folks will also reply. Yours, Bill Tivol
Now that our TEM is fully functional I was hoping to be able to *do* something with it. During sample preparations we have typically used wax/heater to place samples to glass slides, etc. I would rather have an epoxy (acetone soluble) which would eliminate the need for heating during initial polishing. Any suggestions?
Also, we are looking for an epoxy (acetone INSOLUBLE) which can be used for the final specimen, for instance (especially), bonding two silicon wafers back to back before/after milling. Any suggestions here are greatly welcome. We aren't too picky about (non)conducting, just an epoxy which will be strong, adhere very well to silicon to a reasonable temperature range and not lose cohesion in acetone/ethanol, etc.
__ _-==-=_,-. /--`' \_-at---at-.-- { Tim (TJ) LaFave Jr. `--'\ \ {___/. Department of Physics \ \\ " / University of North Carolina, Charlotte } =\\_/` { Charlotte, NC 28223 ____ /= | \_|/ _' `\ _/=== \___/ (704)547-3392 [x4] `___/ //\./=/~\====\ \ // / | ===: http://www.iit.edu/~lafatim | ._/_,__|_ ==: __ \/ \\ \\`--| / \\ ---------- +*+ ---------- | _ \\: /==:-\ `.__' `-____/ |--|==: Such that the future be theirs \ \ ===\ :==:`-' to shape and direct. _} \ ===\ /==/ -----------------------------------------------------------------
Actually, purely from a basic physics point of view an LM is limited to the wavelength of the light used. YOu say you have a 540(nm) light source, which would be an arc lamp or some other source like a laser in order to acheive 540nm rather than a broad range of wavelength values. Especially since you have reported 0.4microns (400nm) as the smallest level of resolution. YOu would need to have 0.4micron (400nm, blue) light to truly be able to have resolution. The lower part of the *visible* range is around 0.25-0.3microns. Hence LM's are limited. EM's on the other hand make use of much smaller wavelengths (higher frequncies) of light to resolve far smaller images...down to angstroms (the size of atoms) (by higher frequencies it is meant that we have higher energies---E = hf.)
So yes it is a silly question, but even as a grad student I think it's good to "recall" these things. Sort of like picking up a history book.
__ _-==-=_,-. /--`' \_-at---at-.-- { Tim (TJ) LaFave Jr. `--'\ \ {___/. Department of Physics \ \\ " / University of North Carolina, Charlotte } =\\_/` { Charlotte, NC 28223 ____ /= | \_|/ _' `\ _/=== \___/ (704)547-3392 [x4] `___/ //\./=/~\====\ \ // / | ===: http://www.iit.edu/~lafatim | ._/_,__|_ ==: __ \/ \\ \\`--| / \\ ---------- +*+ ---------- | _ \\: /==:-\ `.__' `-____/ |--|==: Such that the future be theirs \ \ ===\ :==:`-' to shape and direct. _} \ ===\ /==/ -----------------------------------------------------------------
On Wed, 23 Sep 1998, Ingram, Mike 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 may be a silly question. What is the smallest object ( using } brightfield illumination) that can be observed. For example, I } calculated resolution of .4 micron for a 100x, .90 NA objective and } wavelengh value of 540. This is the smallest separation between } features that can be resolved. Is this also the smallest feature that } can be observed given a single feature on a smooth background? } } Michael Ingram } Rodel, Inc } Polishing Lab } 451 Bellevue Rd } Newark, DE 19713 } (302) 366-0500, ext.: 2545 }
Tim (TJ) LaFave Jr. wrote: ================================================ Also, we are looking for an epoxy (acetone INSOLUBLE) which can be used for the final specimen, for instance (especially), bonding two silicon wafers back to back before/after milling. Any suggestions here are greatly welcome. We aren't too picky about (non)conducting, just an epoxy which will be strong, adhere very well to silicon to a reasonable temperature range and not lose cohesion in acetone/ethanol, etc. ================================================= For this purpose, we have sold quite a bit of an interesting epoxy-phenolic adhesive called M-Bond™ 610 for this purpose. Technical information, including mixing and other use instructions can be found on our website 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 ############################ ==================================================
The best material to use as an acetone soluble mounting material that you=
do not need to heat is a cyanoacrylate material (ie super glue). There have been many suggestions over the years as to the best including Sally Hansen's quick bonding nail glue, Loctite 460, and others. The Loctite product is available from a Loctite distributor and seems to be a good choice especially for Tripod Polishing. For less critical applications, any cyanoacrylate will do. They all vary in viscosity and curing time, b= ut generally if you get a good fresh product it will work. A good acetone soluble low temperature mounting wax is our QuickStick 135. This is a clear wax that flows at about 135 degrees C. The same material is also sold under the name Crystalbond.
As for non-acetone soluble epoxies, the 2 part epoxy that you want is mad= e by:
Epoxy Technology, Inc. 14 Fortune Drive Billerica, MA 01821
TEL: 800-227-2201 FAX: 978-663-9782
The product you want is EpoTek 353ND. They sell it in an 8 ounce kit whi= ch is the smallest size they offer. This is often used in TEM cross section= s.
Another material that is often used is M-Bond 610. That material is made=
} } } } } Please visit us at http://www.southbaytech.com { { { { {
Manufacturers of precision sample preparation equipment and supplies for metallography, crystallography and electron microscopy.
Message text written by "Tim P. LaFave Jr." }
Now that our TEM is fully functional I was hoping to be able to *do* something with it. During sample preparations we have typically used wax/heater to place samples to glass slides, etc. I would rather have an epoxy (acetone soluble) which would eliminate the need for heating during= =
initial polishing. Any suggestions?
Also, we are looking for an epoxy (acetone INSOLUBLE) which can be used for the final specimen, for instance (especially), bonding two silicon wafers back to back before/after milling. Any suggestions here are greatl= y welcome. We aren't too picky about (non)conducting, just an epoxy which will be strong, adhere very well to silicon to a reasonable temperature range and=
not lose cohesion in acetone/ethanol, etc.
__ _-=3D=3D-=3D_,-. /--`' \_-at---at-.-- { Tim (TJ) LaFave Jr. `--'\ \ {___/. =
Department of Physics \ \\ " / =
University of North Carolina, Charlotte } =3D\\_/` { Charlotte, NC 28223 ____ /=3D | \_|/ _' `\ _/=3D=3D=3D \___/ (704)547-3392 [x4] `___/ //\./=3D/~\=3D=3D=3D=3D\ \ // / | =3D=3D=3D: http://www.iit.edu/~lafatim | ._/_,__|_ =3D=3D: __ \/ \\ \\`--| / \\ ---------- +*+ ---------- | _ \\: /=3D=3D:-\ `.__' `-____/ |--|=3D=3D: Such that the future be theirs \ \ =3D=3D=3D\ :=3D=3D:`-= ' to shape and direct. _} \ =3D=3D=3D\ /=3D=3D/ -----------------------------------------------------------------
Thank You for all the answers given to my question about using Helium gas in the ESEM: I asked:
Stowe and Robinson report about reducing beam scattering in conventional Low Vacuum SEM's (Scanning, Vol. 20, 57-60). Are there any experiences using Helium in an ESEM from Electroscan or Philips with a special ESEM-detector? Is the ionization efficiency high enough to get a good performance for amplifying the electrons coming from the sample? How is the image quality compared to e.g. water vapor?
Kind regards
Rainer Ziel ------------------------------------------------------------------------ -------------------------- Roger Moretz wrote:
I can't give you the exact answers you are looking for, but hopefully the following can point you in the right direction. At the '99 MSA meeting in Atlanta, there was a session on ESEM. Several papers were given by the group from the Cavendish Laboratory, especially one on SEM at freezer temperatures by A.L. Fletcher, ... & A.M. McDonald. In an attempt to maintain the proper humidity without icing or thawing, this group used other gases, and determined that N2 was minimally acceptable in terms of obtaining desired imaging SEs. Discussion following the paper included comments on the scattering and SE generation that could be expected from lower atomic number gases, including He. I would suggest your contacting this group. Additionally, I would recommend that you do a literature search for the work of K. Rudiger-Peters and the various publications he has on the physical characteristics and SE generation in the ESEM. I think most of those papers were in the journal Scanning. The principal author on the ESEM is of course Danilatos, and his publications span from 1982 to 1990. Since I do not have ready access to those publications (everything is in boxes, no longer arranged as per my reference manager database) I can't give you figures or exact references even, but I believe that he did publish on different gases. Hope this helps.
Roger Moretz Toxicology Boehringer Ingelheim Pharmaceuticals, Inc. ------------------------------------------------------------------------ ------------------------------------
Bradley L. Thiel wrote:
The question that you posted to the microscopy list server was passed on to me. We have investigated the imaging and amplification properties of several gases in the ESEM. Unfortunately, we have not been very efficient at publishing the results....
Helium actually works very well in the Philips-ElectroScan instruments. However, it is not a very efficient signal amplifying gas, so the emperical results can be a bit deceiving. First, because of the small elastic scattering, the probe-skirt formation is much reduced relative to water vapour. Second, the gas is even less efficient at amplifying the spurious signal from primary beam and backscattered electron ionisations. This means that although the signal collected by the ESD/GSED is not amplified much, it is a very pure secondary signal. The reduced skirt also gives stronger useful contrast.
Unlike other gases, He does not exhibit a peak in its amplification efficiency within the pressure range accessible in the ESEM. It is difficult maintaining pressures above about 8 Torr, because of backstreaming up the column. RP3 has a difficult time pumping the large volume of He.
Consequently, you should use a moderate pressure of gas, avoid detector biases that give arcing, and use the electronic signal amplifier and/or image integration to get a good image.
You may wish to see A.L. Fletcher, B.L. Thiel, and A.M. Donald, Journal of Physics D: Applied Physics, Vol. 30, 2249-2257 (1997). for some discussion of gases.
I hope this is of some help to you. Please feel free to contact me if you have other questions.
Best Wishes, Brad Thiel Polymers & Colloids Group Cavendish Laboratory Department of Physics University of Cambridge ------------------------------------------------------------------------ -------------------- Warren Straszheim wrote:
We routinely use He in our Hitachi 2460N, but we normally run at 40 Pa and use a backscattered electron detector. The resolution is _much_ better than air, and probably better than water vapor. I don't know how it would behave with the specialized sec. e detectors.
Also recall seeing at least one slide on the effect of gas type and pressure on scattering during ESEM sessions at the Atlanta MSA meeting. Sorry, I cannot remember who showed it.
Warren
------------------------------------------------------------------------ -------- Thank You
Rainer Ziel ------------------------------------------------------------- Dipl.-Phys. Rainer Ziel Akzo Nobel Central Research ACR-O/RMG-EM 63784 Obernburg
For an acetone soluble glue that's stronger/better than wax you need to use a cyanoacrylate; any of the family of superglues available in hardware stores. Look on the back of the tube. We use Ross Super Glue and Loctite 460 (preferred). Remember to get a cyanoacrylate *not* an epoxy. Time to dissolve in acetone isn't much different than wax.
For a glue that won't dissolve in acetone we use M-Bond 610 from Measurements Group, Inc., Raleigh, NC ph 919-365-3800, or obtainable from e.m. suppliers. This product is used to bond strain gauges to boilers, etc., but nothing beats it for TEM prep. Er, ah, if you call Measurements Group you may want to concoct a story about having a boiler that needs a strain gauge--better that than see the price go up if they tumble to the fact that the stuff is so good for TEM. :-) Despite their instructions to cure at over 150C for hours we routinely cure it at 70C for less than an hour. No problem at 30C, or even room temp, over night. But isn't 30C = room temp in NC?...never mind.
Ron Anderson, IBM, Hopewell Jct., New York, USA. anderron-at-us.ibm.com IBM Analytical Services; http://www.chips.ibm.com/services/asg
This is a multi-part message in MIME format. --------------B9270C0DCC3F8B822F5E5C45 Content-Type: text/plain; charset=us-ascii Content-Transfer-Encoding: 7bit
Dear Tim,
If you wish to use a product which requires no heat for curing, try the LocTite 460. Depending on the amount of glue placed on the surface, it will harden in about 10 to 15 minutes. Larger mass will require longer cure times.
Every thin film epoxy, will require heating. Epoxies require mass to generate exotherm to cure and a thin film of epoxy does not generate enough heat to cure at RT. Even waxes will require heat, however if you mix the wax with acetone and let it stand, the acetone will evaporate and the wax will cure when this occurs. Time is dependent on the ratio of wax to acetone. M-Bond 610 also requires heat making it useless for your requirement.
We stock both LocTite 460, MBond 610 and EpoxyBond 110(similar to the Epoxy Technology epoxy). Should you have any further questions, please contact me at 800-675-1118 or via email.
Good Luck,
Gary Liechty Allied High Tech Products, Inc.
Tim P. LaFave Jr. 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 } -----------------------------------------------------------------------. } } Now that our TEM is fully functional I was hoping to be able to *do* } something with it. During sample preparations we have typically used } wax/heater to place samples to glass slides, etc. I would rather have an } epoxy (acetone soluble) which would eliminate the need for heating during } initial polishing. Any } suggestions? } } Also, we are looking for an epoxy (acetone INSOLUBLE) which can be used } for the final specimen, for instance (especially), bonding two silicon } wafers back to back before/after milling. Any suggestions here are greatly } welcome. We } aren't too picky about (non)conducting, just an epoxy which will be } strong, adhere very well to silicon to a reasonable temperature range and } not lose cohesion in acetone/ethanol, etc. } } __ _-==-=_,-. } /--`' \_-at---at-.-- { } Tim (TJ) LaFave Jr. `--'\ \ {___/. } Department of Physics \ \\ " / } University of North Carolina, Charlotte } =\\_/` { } Charlotte, NC 28223 ____ /= | \_|/ } _' `\ _/=== \___/ } (704)547-3392 [x4] `___/ //\./=/~\====\ } \ // / | ===: } http://www.iit.edu/~lafatim | ._/_,__|_ ==: __ } \/ \\ \\`--| / \\ } ---------- +*+ ---------- | _ \\: /==:-\ } `.__' `-____/ |--|==: } Such that the future be theirs \ \ ===\ :==:`-' } to shape and direct. _} \ ===\ /==/ } -----------------------------------------------------------------
--------------B9270C0DCC3F8B822F5E5C45 Content-Type: text/x-vcard; charset=us-ascii; name="vcard.vcf" Content-Transfer-Encoding: 7bit Content-Description: Card for Gary Liechty Content-Disposition: attachment; filename="vcard.vcf"
I use M-Bond 610, it comes in an adhesive kit form M-Line Accessories, Measurement Group Inc. Raleigh, NC. M-Bond is designed to mount strain gauges, say to evaluate stress and strain in airplane crashes. The epoxy is a two part, mixed, stored refrigerated, it has about a one month shelf life. Unmixed the two components refrigerated, have a shelf life of about a year. The mix is very thin, low centipoise, due to a high solvent content, this also allows the mix to move into very small spaces, via capillarity.
-- Respectfully, Bob ( Robert G. ) Lawrence Failure Analyst Motorola Phoenix Corporate Research Lab 2100 E. Elliot Rd. MD EL-703 Tempe, AZ 85284-1806 Phone: 602-413-5848 Fax: 602-413-4952 Pager: 1-800-759-7243 PIN 834-2458
We are seeking information on optics that could image the inner surface= of a circular object when shot from above. For example, what type of mirro= r could be placed in the inside of a waste-paper basket to photograph its inner= walls it from above? Our interest is in the simpliest type of mirror (e.g. hemispherical?) with the minimum moving parts.
Please reply off line to: jaincavo-at-goodyear.com Thank you for your help in this matter. =
As far as the acetone insoluble material, I used the LX112 resin kit from Fullam for 7 years (on Si thin sections) with great success. It is now called EPOX 812. Our ratios were as follows: LX112 resin = 5 grams NMA = 4.3 grams DMP - 30 = 7 drops This was well mixed and then the Si slices were "dunked" in the mixture, placed in a teflon vice and then baked at 85 degrees C for 12 hours. This results in a laminate that is resistant to just about everything. It is not as quick as some of the other materials but it is very reliable. Hope this helps.
John Staman Consulting FA Engineer LSI Logic, Colorado Springs
} -----Original Message----- } From: Tim P. LaFave Jr. [SMTP:lafatim-at-charlie.cns.iit.edu] } Sent: Wednesday, September 23, 1998 4:26 PM } To: microscopy-at-sparc5.microscopy.com } Subject: Specimen epoxies for TEM } } ---------------------------------------------------------------------- } -- } 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 } ---------------------------------------------------------------------- } -. } } } Now that our TEM is fully functional I was hoping to be able to *do* } something with it. During sample preparations we have typically used } wax/heater to place samples to glass slides, etc. I would rather have } an } epoxy (acetone soluble) which would eliminate the need for heating } during } initial polishing. Any } suggestions? } } Also, we are looking for an epoxy (acetone INSOLUBLE) which can be } used } for the final specimen, for instance (especially), bonding two silicon } wafers back to back before/after milling. Any suggestions here are } greatly } welcome. We } aren't too picky about (non)conducting, just an epoxy which will be } strong, adhere very well to silicon to a reasonable temperature range } and } not lose cohesion in acetone/ethanol, etc. } } } } __ _-==-=_,-. } /--`' \_-at---at-.-- { } Tim (TJ) LaFave Jr. `--'\ \ {___/. } Department of Physics \ \\ " / } University of North Carolina, Charlotte } =\\_/` { } Charlotte, NC 28223 ____ /= | \_|/ } _' `\ _/=== \___/ } (704)547-3392 [x4] `___/ //\./=/~\====\ } \ // / | ===: } http://www.iit.edu/~lafatim | ._/_,__|_ ==: __ } \/ \\ \\`--| / \\ } ---------- +*+ ---------- | _ \\: /==:-\ } `.__' `-____/ |--|==: } Such that the future be theirs \ \ ===\ :==:`-' } to shape and direct. _} \ ===\ /==/ } ----------------------------------------------------------------- }
We just look delivery of a new Philips XL30/TMP so the ETECs must go. Here's the deal. Three ETEC Autoscans. One is a split console/microscope flavor. One was working, one was semi-working. Two WDS detectors and WDS electronics. You pickup and take away (hint, need a lift gate truck). If no one wants these fine examples of american engineering from the '70 they will be scraped.
Scott
----------------------------------------------------------------------- Scott D. Davilla Phone: 919 489-1757 (tel) 4pi Analysis, Inc. Fax: 919 489-1487 (fax) 3500 Westgate Drive, Suite 403 email: davilla-at-4pi.com Durham, North Carolina 27707-2534 web: http://www.4pi.com
It is widely printed in every textbook to prepare gelatin subbed slides with an addition of chrom alum. Its oxidative properties are supposed to keep the gelatin from taking on bacterial populations. Bacteria are no problem if the gelatin solution is used, and discarded. However, there is a lot of grief with fading of sections, or color change of sections, when stained with the common LM stains. Fading of sections is a complicated problem, but one factor is the propensity of LM stains to react to redox shifts. Therefore, do not use chrome alum in subbing solutions. What if you use the slides for immunostaining and you have a nice underlayer of chrome alum below the section? Chrome alum makes perfect sense in the path lab perhaps. That is where it started - with paraffin sections. Many, many conceptual errors have been made by simply applying the methodologies designed for paraffin section to methods employed in TEM. Those two research specialties have very little in common. Hildy
Maleate buffers are not used for primary or secondary fixation. Why not use cacadylate? The maleate buffer system is very useful for enbloc use of UA in tissues which have been previously treated with osmium. They can be made to approximate the pH of UA in solution, causing the osmium to be stable and not move slightly. (this method can yield most beautiful membrances) If you want a reference on that, let me know. Why not use cacadylate buffer? It is toxic, but it would seem to be the method of choice if your specimens tolerate it. Make stock cocadylate at two times the strength. Wear a mask when weighing it. The powder is dangerous. Bye, Hildy
Does anyone know where Rapidoprint DD37E repair parts and chemicals might be obtained? Also, does anyone know if Kodak still makes Motion Picture #5302 is still made? TIA.
Chuck Butterick Engineered Carbons, Inc. Borger, Texas
Can anyone tell me a good way of removing the polymer packaging from an IC chip? I'd like to strip away the packaging and use the IC as an SEM specimen.
Many thanks, steve
-- Steven Kim | stevekim-at-nwu.edu MLSF, Room 3078 | 847.491.5888 (work) 2225 N. Campus Dr. | 847.491.7820 (fax) Evanston, IL 60208 | 847.332.1069 (home)
Dear All, In my reply to the Listserver of 9/17/98 on this subject, I mistakenly stated that Link had warmed the snout of their detector to prevent oil from settling on the detector. This, in fact, was the work of one individual on his Link detector, and did not work. The message I got from Link, privately, is as follows:
You recently sent a message to the microscopy listserver on the subject of oil contamination. The text was as follows:- ************* This is a problem of all SEMs, not just JEOL, and really is a result of the EDS detector being the coolest spot in the chamber. The cleanliness of the vacuum system just regulates how long the contamination will take to build up. Link solved this problem by warming the snout of their detector. This doesn't solve the problem of oil contamination, it just moves it away from the EDS detector. I agree woth Steve Chapman that the oil is from the rotary pump. *************
I should just like to correct a few points in this message.
The conditioning circuit fitted to the Oxford Link detectors DOES NOT warm the snout of the detector. The conditioner acts on the internal components of the detector, and cannot remove oil contamination. Warming the end of an oil contaminated detector by any means is likely to cause expensive damage to the window.
If the end of a detector becomes contaminated with oil, cleaning must only be attempted by a qualified engineer - the windows are fragile, and if cleaning is attempted in the wrong way the window could fail.
We would appreciate it if you could publish a note correcting these points, using your own words - we don't want people reporting window failures because they have been warming the ends of their detectors to remove oil!
Thanks
Chris Lay Customer Support Manager Oxford Instruments Microanalysis Group
*********************************************** Microanalysis Group, Oxford Instruments Halifax Road, High Wycombe Buckinghamshire, HP12 3SE, England Tel: +44 (0)1494 442255 Fax: +44 (0)1494 524129 URL http://www.oxford-instruments.com/ *********************************************** Needless to say, I was somewhat taken aback by the harsh tone of this message, and I was suprised to learn that customers cannot clean the window of a Link detector themselves. My experience with a Kevex Quantum window detector is that serious work with light elements must be preceeded by a careful solvent wash of the window, since the thinnest oil film will seriously absorb the lighter elements. Kevex have a Technical Bulletin (#21) showing this very graffically on SiO2, before and after cleaning, and complete instructions on cleaning. I always clean the detector, now, just before anyone does light element work. I have a Link detector, but a service call from a qualified Link engineer would cost airfare for 3000 miles plus time charges and supplies: a minimum of $3500 CAN. Sorry if I misled anyone.
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 fax: 604-822-3619 e-mail: mager-at-interchg.ubc.ca
I have reacted epoxy-like material by slowly dropping boiling, red fuming nitric acid on the chip. Be careful! This acid is very reactive and will produce noxious/toxic fumes.
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Can anyone tell me a good way of removing the polymer packaging from an IC chip? I'd like to strip away the packaging and use the IC as an SEM specimen.
Many thanks, steve
-- Steven Kim | stevekim-at-nwu.edu MLSF, Room 3078 | 847.491.5888 (work) 2225 N. Campus Dr. | 847.491.7820 (fax) Evanston, IL 60208 | 847.332.1069 (home)
via smtpd (for sparc5.microscopy.com [206.69.208.10]) with SMTP; 25 Sep 1998 13:20:16 UT Received: from ptag2 (ptag2.pt.cyanamid.com [141.173.51.2]) by igate.Cyanamid.COM (8.8.6/8.8.6) with ESMTP id JAA25089 for {Microscopy-at-MSA.Microscopy.com} ; Fri, 25 Sep 1998 09:19:54 -0400 (EDT) Received: from ccmail.pt.Cyanamid.COM by pt.Cyanamid.COM (PMDF V5.1-9 #28913) id {01J27VI3Y90W8X1M0W-at-pt.Cyanamid.COM} for Microscopy-at-MSA.Microscopy.com; Fri, 25 Sep 1998 09:20:09 EDT
For those of us in other fields with some curiosity, what is this word? I assume it's short for something. Is it just glue or does it have magical properties?
Leonard Corwin Research Chemist Fort Dodge Animal Health Princeton, NJ 08543-0400
I need the manual for an older AO Interference scope. There is no model number on the unit, but I believe it is Series 7 or 9, and I believe the manual is:
I would appreciate some input on how to successfully perform antigen unmasking on formaldehyde fixed archived skin. I have experimented = with 3 commercial compounds and had only moderate success. Most of the problems seems to stem from the multiple layer nature of skin and the thickness of the tissue. I would ideally like to be able to perform this procedure on section upward of 25 =B5m and up to 100 =B5m. At = this thickness the tissue has a hard enough staying on the slide w/o being microwaved or steamed. I have tried the same compounds + protocols = with free floating sections, but the results were equally unimpressive.
I do realize that I may be forced to cut much thinner sections, = possibly around 10=B5m. Any input on successful unmasking procedures and the thickness of the tissue the procedure was performed on would be much appreciated.
Ramin Rahbari PARKE-DAVIS Pharmaceutical Research Pathology and Experimental Toxicology 2800 Plymouth Road Ann Arbor, MI 48105 Office (734) 622-3383 Fax (734) 622-5001 Ramin.Rahbari-at-WL.COM
If this is the typical "phenolic", black resin plastic, fuming nitric acid brought to approximately 90 degrees C will do the job. My suggestion is to put the part in some type of "basket" to facilitate easy removal and progress monitoring. Depending on the size of the part, it usually takes 5 - 8 minutes (most of the time) at that temperature.
John Staman Consulting FA Engineer LSI Logic, Colorado Springs
} -----Original Message----- } From: Steven Kim [SMTP:stevekim-at-casbah.acns.nwu.edu] } Sent: Thursday, September 24, 1998 4:38 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Solvent for IC Packaging? } } ---------------------------------------------------------------------- } -- } 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 } ---------------------------------------------------------------------- } -. } } } Can anyone tell me a good way of removing the polymer packaging from } an IC } chip? I'd like to strip away the packaging and use the IC as an SEM } specimen. } } Many thanks, } steve } } -- } Steven Kim | stevekim-at-nwu.edu } MLSF, Room 3078 | 847.491.5888 (work) } 2225 N. Campus Dr. | 847.491.7820 (fax) } Evanston, IL 60208 | 847.332.1069 (home) } } http://pubweb.acns.nwu.edu/~stevekim } }
This is a multi-part message in MIME format. --------------A6FDFD211AC81FB3B5409A39 Content-Type: text/plain; charset=us-ascii Content-Transfer-Encoding: 7bit
Hello Steven,
There is a company in Hayward Ca, called EKC Technology that sells a product you are looking for. It is called EKC 270 and will remove the px from the device under heat in about 30 to 45 minutes, depending on the thickness.
Good Luck,
Gary Liechty Allied High Tech Products, Inc.
Steven Kim 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 } -----------------------------------------------------------------------. } } Can anyone tell me a good way of removing the polymer packaging from an IC } chip? I'd like to strip away the packaging and use the IC as an SEM } specimen. } } Many thanks, } steve } } -- } Steven Kim | stevekim-at-nwu.edu } MLSF, Room 3078 | 847.491.5888 (work) } 2225 N. Campus Dr. | 847.491.7820 (fax) } Evanston, IL 60208 | 847.332.1069 (home) } } http://pubweb.acns.nwu.edu/~stevekim
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I have used boiling sulfuric acid to remove epoxy packaging. I usually cut off all the metal leads first. It can take all day depending on the thickness, but this has the advantage of not removing the aluminum traces (some of the time!)
best regards mark
Mark W. Lund, PhD Director } } Soft X-ray Web page http://www.moxtek.com { { MOXTEK, Inc. 452 West 1260 North Orem UT 84057 801-225-0930 FAX 801-221-1121 lundm-at-xray.byu.edu
"The state is good at simple tasks, like killing people and seizing their wealth. It has far more trouble reaching inside individuals and making them good." Doug Bandow
I'm interested in making/buying an amplifier/pre-amp for specimen current measurements. Does anyone use a Stanford Research Systems SR570? If so, how do you like it? Any suggestions for getting the best performance from SC as an imaging technique?
Thanks in advance,
Harold J. Crossman OSRAM SYLVANIA INC. Lighting Research Center 71 Cherry Hill Dr. Beverly, MA 01915 (978) 750-1717 crossman-at-osi.sylvania.com
} go. Here's the deal. Three ETEC Autoscans. One is a split } console/microscope flavor. One was working, one was semi-working. Two WDS } detectors and WDS electronics. You pickup and take away (hint, need a lift } gate truck).
The ETECs have been taken.
Scott ----------------------------------------------------------------------- Scott D. Davilla Phone: 919 489-1757 (tel) 4pi Analysis, Inc. Fax: 919 489-1487 (fax) 3500 Westgate Drive, Suite 403 email: davilla-at-4pi.com Durham, North Carolina 27707-2534 web: http://www.4pi.com
I want to thank all of you who take the time to help me out when I am up a creek, or about to embark on something retarded. You have saved me endless hours of grief with your sharing of knowledge and experience. In return I try to help where I can. (The refrences you want for the maleate enbloc UA I will find on Tuesday). Bye, Hildy
A few more caveats and questions - helium can certainly produce a dramatic difference in image quality and the amount of X-ray scatter, especially at lower kVs. We are using it a lot in CL imaging as well. However there are a couple of complications arising from its increased thermal conductivity relative to nitrogen or air. -
- A liquid nitrogen-cooled stage requires noticably more LN2 - so temperature at the specimen surface MAY be a little higher than in other gases, depending on the detector positions I suppose.
More importantly -2. Pressure readout from a gauge depending on a thermal effect, as in the Piranis used in most chamber pressure control systems, will be more in helium than nitrogen at the same pressure. There are graphs around. The diifference is not very much in the 0.01-0.1 torr range, but diverges very rapidly - from memory an indicated pressure of 3.5-4 torr corresponds to a true pressure of only about 1.3 torr. Which may affect the pressure range a gaseous amplification system thinks it is working at? (Ionisation potential of helium is about 25eV, cf 15eV for nitrogen...I dont know if that is enough to have a big effect on detector efficiency?) We only use BSE and CL, so I've no data on this, but for what its worth in the pressure range we use, we dont get the swamping background CL signal in helium that comes in at higher pressures in air.
3. Permeability -the EDS on our VP system has a Be window, and we havent had obvious problems resulting from helium. However somebody with an ultra-thin window EDS once mentioned that after using helium for leak testing, they thought some gas had found its way into the EDS detector. Not a good prospect, especially given the thermal properties. Does anyone have any more information or experience relating to this?
Sally Stowe
} Rainer Ziel wrote - } Thank You for all the answers given to my question about using Helium } gas in the ESEM: } I asked: } } Stowe and Robinson report about reducing beam scattering in conventional } Low Vacuum SEM's (Scanning, Vol. 20, 57-60). Are there any experiences } using Helium in an ESEM from Electroscan or Philips with a special } ESEM-detector? Is the ionization efficiency high enough to get a good } performance for amplifying the electrons coming from the sample? How is } the image quality compared to e.g. water vapor? } } Kind regards } } Rainer Ziel } ------------------------------------------------------------------------ } -------------------------- } Roger Moretz wrote: } } I can't give you the exact answers you are looking for, but hopefully } the following can point you in the right direction. At the '99 MSA } meeting in Atlanta, there was a session on ESEM. Several papers were } given by the group from the Cavendish Laboratory, especially one on SEM } at freezer temperatures by A.L. Fletcher, ... & A.M. McDonald. In an } attempt to maintain the proper humidity without icing or thawing, this } group used other gases, and determined that N2 was minimally acceptable } in terms of obtaining desired imaging SEs. Discussion following the } paper included comments on the scattering and SE generation that could } be expected from lower atomic number gases, including He. I would } suggest your contacting this group. Additionally, I would recommend } that you do a literature search for the work of K. Rudiger-Peters and } the various publications he has on the physical characteristics and SE } generation in the ESEM. I think most of those papers were in the } journal Scanning. The principal author on the ESEM is of course } Danilatos, and his publications span from 1982 to 1990. Since I do not } have ready access to those publications (everything is in boxes, no } longer arranged as per my reference manager database) I can't give you } figures or exact references even, but I believe that he did publish on } different gases. Hope this helps. } } Roger Moretz } Toxicology } Boehringer Ingelheim Pharmaceuticals, Inc. } ------------------------------------------------------------------------ } ------------------------------------ } } Bradley L. Thiel wrote: } } The question that you posted to the microscopy list server was passed on } to me. We have investigated the imaging and amplification properties of } several gases in the ESEM. Unfortunately, we have not been very } efficient } at publishing the results.... } } Helium actually works very well in the Philips-ElectroScan instruments. } However, it is not a very efficient signal amplifying gas, so the } emperical results can be a bit deceiving. First, because of the small } elastic scattering, the probe-skirt formation is much reduced relative } to } water vapour. Second, the gas is even less efficient at amplifying the } spurious signal from primary beam and backscattered electron } ionisations. } This means that although the signal collected by the ESD/GSED is not } amplified much, it is a very pure secondary signal. The reduced skirt } also gives stronger useful contrast. } } Unlike other gases, He does not exhibit a peak in its amplification } efficiency within the pressure range accessible in the ESEM. It is } difficult maintaining pressures above about 8 Torr, because of } backstreaming up the column. RP3 has a difficult time pumping the large } volume of He. } } Consequently, you should use a moderate pressure of gas, avoid detector } biases that give arcing, and use the electronic signal amplifier and/or } image integration to get a good image. } } You may wish to see } A.L. Fletcher, B.L. Thiel, and A.M. Donald, Journal of Physics D: } Applied } Physics, Vol. 30, 2249-2257 (1997). } for some discussion of gases. } } I hope this is of some help to you. Please feel free to contact me if } you } have other questions. } } Best Wishes, } Brad Thiel } Polymers & Colloids Group } Cavendish Laboratory } Department of Physics } University of Cambridge } ------------------------------------------------------------------------ } -------------------- } Warren Straszheim wrote: } } We routinely use He in our Hitachi 2460N, but we normally run at 40 Pa } and } use a backscattered electron detector. The resolution is _much_ better } than } air, and probably better than water vapor. I don't know how it would } behave } with the specialized sec. e detectors. } } Also recall seeing at least one slide on the effect of gas type and } pressure } on scattering during ESEM sessions at the Atlanta MSA meeting. Sorry, I } cannot remember who showed it. } } Warren } } ------------------------------------------------------------------------ } -------- } Thank You } } Rainer Ziel } ------------------------------------------------------------- } Dipl.-Phys. Rainer Ziel } Akzo Nobel Central Research } ACR-O/RMG-EM } 63784 Obernburg } } Tel: (06022) 81-2645 } Fax: (06022) 81-2896 } E-mail: Rainer.Ziel-at-AkzoNobel.com } } } } ---------------------------------------------------------------------- Dr Sally Stowe |Email: stowe-at-rsbs.anu.edu.au Facility Coordinator |Post: GPO Box 475 ANU Electron Microscopy Unit |ANUEMU (RSBS) Ph 61 (0)2 6249 2743 |Australian National Univ. FAX 61 (0)2 6249 4891 |Canberra, Australia 2601 http://online.anu.edu.au/EMU/home.htm
A while back (a year ago), I was told by AGFA that the DD3700 was being phased out and would no longer be serviced and that graded Rapitone paper P 1-4, ETC would no longer be available. We now use polycontrast resin coated paper, but most older users are quite unhappy with using the Kodak filter sets to get contrast. As a result, we are switching over to the Mohr-Pro 8 system and also doing more digital imaging. I have four AGFA DD3700's which are about to be surplused, along with many extra parts which I have been stock piling for years. If any one is interested in obtaining these items, let me know. Also, if anyone knows of a source for graded rapitone papers that are still good, let me know and maybe we can work out a deal. Does any one out there like their Mohr-Pro 8 system? Any long term issues, maintenance, paper, ETC to watch out for ? Thanks, Thomas A Baginski, Room G-230 Technical Coordinator for Microscopy Uniformed Services University of the Health Sciences 4301 Jones Bridge Road Bethesda, MD 20814-4799
Postdoctoral Position in Environmental Catalysis/HREM
A postdoctoral position is available at Northwestern University to work on Environmental Catalysis. The work will involve in-situ work using a unique UHV-HREM and gas treatment of various samples (see http://www.numis.nwu.edu for some information about the hardware). A strong background in HREM and crystallography is highly desirable, and experience in catalysis and/or surface science will be significant; extensive experience in other types of TEM are an alternative. To apply, send an email to L. D. Marks at ldm3-at-apollo.numis.nwu.edu including a CV (text only, no attachments) and the names of referees.
++++++++++++++++++++++++++++++++++++++++++++++++ Laurence Marks Department of Materials Science and Engineering Northwestern University fax: (847) 491-7820 mailto:l-marks-at-nwu.edu http://www.numis.nwu.edu ++++++++++++++++++++++++++++++++++++++++++++++++
Dear Sally, } } 3. Permeability -the EDS on our VP system has a Be window, and we } havent had obvious problems resulting from helium. However somebody } with an ultra-thin window EDS once mentioned that after using helium } for leak testing, they thought some gas had found its way into the } EDS detector. Not a good prospect, especially given the thermal } properties. Does anyone have any more information or experience } relating to this? } When I was a grad student studying the scattering of protons on He, we had a target which was an evacuated, completely sealed thin glass sphere. The target was filled by immersing the sphere in a container filled with He. The He would diffuse through the sphere, which we would then use until sufficient He had diffused out. Since the glass was thick enough to be mechanically self-supporting, one can conclude that He will diffuse quite readily through many materials usually considered imper- meable. I'm not surprised that He can get through an ultra-thin window, but I would be surprised if it didn't also get through a Be window. I would also not expect obvious problems from the relatively small amount of He which got through the small area of an EDS window. Yours, Bill Tivol
Hi: If you have a complete Zeiss 109 or 10C for sale or if you know of any please contact me. It does not have to be in working condition. Thank you, Peter Jordan, EMSI 909 694-1839
To prevent tissues from schrinkage, EtOH 50% can be used instead of 70%. This is a very good fixative for general plant anatomy, it can be used as a preservation fluid as well: samples can be stored in it for years. Large specimens can be fixed. Fixation of mitotic figures and nuclear detail is poor... Fixation time for specimens 1cmx1cmx0.5cm: at least 24h. washing in EtOH 70% or 50%, depending on EtOH concentration used in the fixative. _______________
Chrom-acetic acid -------------------------- CrO3, 10% soln in dist. water: 2.5 - 10ml acetic acid (10%): 5 - 10ml add distilled water to obtain 100ml of fixative. Prepare just before use. Keep samples as small as possible. Fixation time: not longer than necessary (max. 24h), otherwise: maceration! Wash in running (or at least frequently changed) water, untill most of the yellow stain is removed (complete removal is difficult!). Fixation and washing in the dark to prevent the formation of chromates. Good preservation of nuclear detail and almost all structures (mitochondria and proplastids are frequently lost). Fixatives containing CrO3 are good mordants prior to staining with safranin. Staining with hematoxylins (f.e. Delafield's) is dfficult, but Fe-hematoxylins (Heidenhain's, Weigert's) can be used. Waste disposal of chrome is problematic: check with your local authorities! _______________
Bouin -------- This wel known fixative for animal histology can also be used for plant tissues.
Prepare just before use (there are conflicting opinions regarding the shelf life of the solution).
Fixation time: up to 3 days. Wash in EtOH 70% untill most of the yellow color is removed.
---------- } Van: Botanique1-at-aol.com-at-sparc5.microscopy.com } Aan: Microscopy-at-sparc5.microscopy.com } Onderwerp: Glandular trichomes } Datum: zondag 27 september 1998 7:01 } } ------------------------------------------------------------------------ } 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'm looking for a fixation recipe for herbaceous plants... I'm wanting to } look at glandular trichomes.... Any comments are welcome! } } Stefanie } ****************** } Stefanie Galgon } Department of Biology } Northern Arizona University } smg4-at-dana.ucc.nau.edu } Botanique1-at-aol.com }
Thank you to those who have replied. I do appreciate the help. However, I realize now, that in my hasty initial email, I left out ALL the details... silly me! Now that I have a moment, I'll say a little more....
For a few years now, I have been looking at the differences of concentrations of particular terpenes in Salvia officinalis. A main focus has been the differences in young and mature leaves. I now want to compare anatomy and morphology of peltate glandular trichomes (the terminology is so scattered in this area... peltate, sessile etc.) in young and mature leaves using TEM. Fun stuff, eh?
During my literature search, I came across quite a few SEM papers and recipes. I figured I would just go with one of the basics I am familiar with, but I thought shooting the question out to you guys wouldn't hurt.
Thanks again for the input.
Regards,
Stefanie ****************** Stefanie Galgon Department of Biology Northern Arizona University smg4-at-dana.ucc.nau.edu Botanique1-at-aol.com
Dear All, Can anybody let me know the recipe for twinning-jet-polish of Ag? Thanks in advance. Ke Han Center for Materials Science Los Alamos National Laborotory Los Alamos, New Mexico NM87545, USA Tel: 505-6650771 Fax: 505-6652992
I know two people looking for employment in EM/microscopy . One has managed an EM lab for many years - their lab is about to close. The other is a post-doc in Genetics with lots of microscopy experience.
Any help/leads would be greatly appreciated.
Beth
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
Ken: The following is for the window technique but it might work for jet polishing: aqueous potassium cyanide at less than 20 degrees C. voltage of about 8 V and current density of 1.8A/cm.sq. The sample must be rinsed in distilled water and used at once or stored in ethanol} Source: Desmond Kay editor. Pg. 403.
Jordi Marti ---------- } From: ke Han To: microscopy-at-sparc5.microscopy.com -----------------------------------------------------------------------
A colleague has an EMITECH K850 CPD unit that appears to be malfunctioning.
1. Can any one supply us with information on a US distributor who might me able to help?
2. Can anyone help us diagnose the problem? The unit is 3 years old but has never been used. Upon presurization with CO2, there is a large gas leak originating from inside the unit. Upon removal of the back, the leak is seen to originate from what I am guessing is the overpressure/ heating override sensor. In other words, the leak is coming from a stainless steel or chromed sensor that has a copper CO2 line in one end and two electrical leads off the top (brown and yellow?--I forget exactly what the colors are). My guess is that this is the sensor that disables the heater when some pre-set pressure is exceeded, but I could be wrong. At any rate, the electrical leads on top of the sensor extend from a black plastic insert--the leak is between the black plastic insert and the SS surround. Does anyone know if this sensor is repairable?
TIA
Bob
Dr. Robert R. Wise Department of Biology and Microbiology University of Wisconsin-Oshkosh Oshkosh, WI 54901
(920) 424-3404 tel (920) 424-1101 fax wise-at-uwosh.edu www.uwosh.edu/departments/biology/wise/wise.html
Details of the October 9, 1998, Midwest Microscopy and Microanalysis Me= eting at Purdue University are now posted on the MSA webpage at http://msa.microscopy.com/. Follow the Local Affiliate Societies links= to find the agenda and maps. For further information, you may contact me = via e-mail. Thanks!
Jane A. Fagerland, Ph.D. Dept. Microscopy and Microanalysis Abbott Laboratories Abbott Park IL 60064 =
{excerpt} {fontfamily} {param} Times {/param} Postdoctoral Position=20
An NIH-funded postdoctoral position is available immediately for structural studies of HIV-1 gp120. This is a joint project between Ken Roux and Ken Taylor.=A0 The project involves three dimensional epitope mapping and segmental flexibility analyses of gp120, gp160 and polymers there of in complex with ligands and monoclonal antibodies, using electron microscopy=A0 (negative staining and cryo-EM) and computational image analysis.=A0 Experience in electron microscopy and/or computational image analysis is essential. The project provides opportunities for development of innovative approaches to single particle microscopy and image analysis.=A0 State of the art microscopy facilities include a Philips CM300-FEG and a Philips CM120, each equipped with a fully computer controlled goniometer (CompuStage) for tomography and low dose cryoEM data collection.=A0 Computational facilities include a cluster of DEC Alpha compute servers and numerous Silicon Graphics workstations.=A0 Digitizing facilities include a Perkin-Elmer PDS 1010M densitometer.=A0 The labs and facilities are part of an extensive interdisciplinary Structural Biology program with expertise in X-ray crystallography, NMR, protein engineering, 3-D electron microscopy.=A0=20
We have a copy of your E mail on the Server, I believe that you have been given the address on our USA and UK Office.
However, from your detailed description your analysis of the problem would appear to be correct. This switches off heaters on an overpressure, which normally does not occur in normal use.
These are the seal units which can't be repaired in a local situation, and Emitech will need to return it to the original manufacturer.
However as part of our Service Support Policy, which gives you two years warranty on all parts, we would be pleased to extend this and send you an advanced replacement free of charge. We would be pleased if you could then return the old one.
Please advise delivery address.
Kind Regards,
David Robinson
In message {v04003a04b23548a2ce8b-at-[141.233.130.134]} , "wise-at-vaxa.cis.uwo sh.edu"-at-Sparc5.Microscopy.Com writes } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} ------------------------------------------------------------------------ } 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, } } We want to integrate a focused ion beam gun into our large chamber SEM. Who } has experience } in doing this? } } Thanks in advance, Martin Klein } ___________________________ } } VisiTec Microtechnik GmbH } Karl-Marx-Str. 14 } D-23936 Grevesmuehlen } Germany } } Fon: +49-3881-790-49 } Fax: +49-3881-790-48 } email: mklein-at-visitec-em.de } web site: www.visitec-em.de } } } }
Please help me out on this, as its been many years since I have used permount and similar mterials.
A user is mounting ostracods in Cytoseal. Initially, she has beautiful mounts. After a few days of drying at room temperature, however, she finds lots of bubbles under her coverslip. Is this buffer/solvent leaching out of her sample? What can she do to solve this problem?
thanks in advance
steve
--------------------------------------------------------------------- Dr. Steven Barlow, Associate Director EM Facility/Biology Department San Diego State University 5500 Campanile Drive San Diego CA 92182-4614 phone: (619)594-4523 fax: (619) 594-8759 email: sbarlow-at-sunstroke.sdsu.edu website: http://www.sci.sdsu.edu/emfacility/
JEOL Geremany. Dr. W. Knoll 0049/8165-77346 0049/8165-77512 (fax)
-----Original Message----- } From: Dr P. Echlin {pe13-at-cus.cam.ac.uk} To: Martin Klein {Visitec-at-t-online.de} Cc: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com}
We are looking for image analysis software that will detect small area= s of color (say yellow on black) and automatically send a relay to a control= system when a colored area is detected? Thanks in advance for your attention = to this posting. Please reply off line to: jaincavo-at-goodyear.com =
Has anyone had experience in resolving Golgi with 100x fluroescence microscopy?
I am trying to discern the tubular-vesicular mechanisms of PC-12 cells through fluorescence microscopy and want to label Golgi with BODIPY ceramide, but fear they are too small to show up decently.
Appriciate the help!
-- Vickie A. Kimler, Ph.D. Assistant Professor of Biology and Allied Health Director, Cancer Research Facility Mercyhurst College 501 East 38th St. Erie, PA 16546 Voice: 814-824-2169 FAX: 814-824-2188 e-mail: {vkimler-at-mercyhurst.edu}
Ke Han asked "Dear All, Can anybody let me know the recipe for twinning-jet-polish of
Ag?"
Bernie Kestel used the following technique on South Bay 550-B single jet instrument with good results: 60 ml perchloric acid 460 ml ethyl alcohol 280 ml butyl alcohol 150 ml butyl cellosolve
Temperature: -20 degrees centigrade. Voltage: 220 V. Current: 70mA. with a thin plastic diaphram having a 1.4 mm center hole retaining the specimen in it's holder. In -situ, magnified viewing of the polishing and the unit's 300 volt capability were very helpful.
-- Dr. Roseann Csencsits Electron Microscopy Center Building 212/C217 Argonne National Laboratory 9700 South Cass Avenue Argonne, IL 60439-4838 Phone: (630) 252-4977 Fax: (630) 252-4798
You can add a drop of xylene to dissolve the mounting medium then expose the slide to a vacuum for a few seconds to remove the bubles. For more details see the above reference.
Jim Harper microfab-at-aol.com
In a message dated 98-09-29 15:38:23 EDT, you write:
{ { Please help me out on this, as its been many years since I have used permount and similar mterials.
A user is mounting ostracods in Cytoseal. Initially, she has beautiful mounts. After a few days of drying at room temperature, however, she finds lots of bubbles under her coverslip. Is this buffer/solvent leaching out of her sample? What can she do to solve this problem?
thanks in advance
steve
--------------------------------------------------------------------- Dr. Steven Barlow, Associate Director EM Facility/Biology Department San Diego State University 5500 Campanile Drive San Diego CA 92182-4614 phone: (619)594-4523 fax: (619) 594-8759 email: sbarlow-at-sunstroke.sdsu.edu website: http://www.sci.sdsu.edu/emfacility/ } }
Discussion came up regarding the use of maleate buffers. Some people asked for a protocol. I will give you the reference, as writing it out would take a long time. This protocol is wonderful for the preservations of membranes, and great for preservation in general. It calls for GA,PAF prefix in phos buffer (or cac, if absolutely necessary), a one second rinse and then postfixation in veronal acetate buffered osmium. Veronal buffer cannot be used as a prefixation buffer, but the bacteriologist have used it for eons for good membrane preservation. I have left the material to fix overnight in osmium, if I had a structure of special interest which contained a lot of unsaturated lipid. Next the prep is washed with maleate of a certain pH in order not to dislocate the osmium. The material is then enbloc stained with UA in maleate. There are no huge shifts of pH here in this method. Acetone dehydration (saves lipids), PO intermediate and epoxy follows. All steps are done on ice except the last 100% acetone. All this was backed up at an EMSA meeting years ago by Janet Boyne of SF. Her talk appeared in the Bulletin.
Here is the reference. Williams MC. Conversion of lamellar body membranes into tubular myelin in alveoli of fetal rat lungs. J Cell Biol 1977; 72:260
If you try this method and like it, make up large quantities of the buffers and freeze them in aliquots. Bye, Hildy
I have inherited an old Ladd Sputter coater, the unit works, but the
film thickness is twice that indicated on the meter. This machine has a
density switch to enter the density of the material being sputtered, it is set to the correct density. I have been working with Ladd and they have been very helpful, but this unit is so old and they have moved several times.
I am looking for a schematic/ manual for a Poloron MK1 Film Thickness Monitor, it is used in a LADD Sputter Coater, model 30800 , the Ladd model number on the film thickness monitor is 30808. Any help would be greatly appreciated.
-- Respectfully, Bob ( Robert G. ) Lawrence Failure Analyst Motorola Phoenix Corporate Research Lab 2100 E. Elliot Rd. MD EL-703 Tempe, AZ 85284-1806 Phone: 602-413-5848 Fax: 602-413-4952 Pager: 1-800-759-7243 PIN 834-2458
Hello everyone; I am about to revisit in situ hybridization using DIG labelled probes. One thing I remember from the course I took was to NOT use Permount to mount the slides at the end of the protocol, or other organic based mounting media. My inclination is to use glycerol, but this doesn't seem to be a "permanent" treatment. Any suggestions would be greatly appreciated.
thanks in advance shea Dr. S. Shea Miller Agriculture & Agri-Food Canada Eastern Cereal & Oilseed Research Centre Rm 2068, Bldg 20, CEF Ottawa, Ontario Canada K1A 0C6 Phone: (613)759-1760 Fax: (613)759-1701 e-mail: millers-at-em.agr.ca
I got no responses from my request of last week, so I am calling out again in the hope that someone on the list does indeed have this manual:
AO-Baker Interference Microscope, a Series 7 or 9 (I believe) built upon a basic AO Series 4 binocular scope, made probably in the late 50s or early 60s
Again, I will pay for copying costs and postage.
ALTERNATIVELY: I heard that someone in Canada has been collecting old microscope manuals for resale. Does anyone know who this person is?
Microscopy List Server {Microscopy-at-Sparc5.Microscopy.Com}
Dear Recipient,
I am hoping to reach email or internet catch-all addresses for microscopy secondhand equipment dealers.
Thieves stole three slide cabinets from our histology teaching lab over the last weekend. We bolted the stable door after the horse had flown, they came back yesterday lunch hour, forced the door, and took three more.
The cabinets must be valuable. The slides that they contained were in part invaluable.
I knew about the confocal list server, but I figure that confocal users will not be the class of person looking for antique slide filing cabinets. One of them was a rare [I guess late Victorian] beauty, mahogany, glass doors and ivory handles on all the 30 drawers. That was, of course, the one with our most valuable teaching slides in. The others were of a white coloured wood, probably beech.
I cannot imagine that the thieves would try to trade the slides because they would be too easily identified.
I would like to circulate to people who might be offerred these goods for sale and to ask them to contact us for further identification.
Yours sincerely,
Alan Boyde
Alan Boyde, Professor Hard Tissue Research Unit Department of Anatomy and Developmental Biology University College London Gower Street London WC1E 6BT United Kingdom Phone: If I do not answer on +44 171 419 3316 [or 3320/3322/3321] There are recording machines for messages up to 3 minutes on 3313 [and 3315] FAX +44 171 391 1302
On Tue, 29 Sep 1998 14:57:40 +0000 "Vickie A. Kimler, Ph.D." {vkimler-at-mercyhurst.edu} wrote:
} } } Has anyone had experience in resolving Golgi with 100x fluroescence } microscopy? } } I am trying to discern the tubular-vesicular mechanisms of PC-12 cells } through fluorescence microscopy and want to label Golgi with BODIPY } ceramide, but fear they are too small to show up decently. } } Appriciate the help! } } Vickie,
Its a piece of cake. We have undergrads do ceremide-Golgi labeling in lab exercises and they get beautiful pictures to take home and wow their parents, when recorded on a color video printer. One wrinkle that helps: Incubate the cells in ceremide for 10' at 4 deg C, then warm to 37. The dye will bind to the cell surfaces at 4 and be taken up synchronously and transferred to the Golgi when the cells are warmed.
-Dennis ---------------------- Dennis Goode Dept. of Biology University of Maryland College Park 20742 dg0-at-umail.umd.edu
} Microscopy List Server {Microscopy-at-Sparc5.Microscopy.Com} } } Dear Recipient, } } I am hoping to reach email or internet catch-all addresses for microscopy } secondhand equipment dealers.
To the list:
We all have sympathy when theft like this happens. Fear and loathing are two thoughts that come to mind.
Perhaps the following story will help a little.
Several years ago someone stole a computer that was a part of an imaging workstation. The poor soul whose lab was hit had his wits about him. He placed a classified ad in a local buy and swap newspaper (one that was known to list stolen merchandise from time to time). The ad read something like: "Desparate, looking for an older computer board with a serial number from between n and n+x. The board is usually mounted in a computer model XX. Will pay top dollar for the board." He gave his home phone number. He got a phone call within 24 hours. He actually got the caller to give him the serial number off of the computer. It was the lab computer. He arranged to go over to the caller's house to buy the board; of course, accompanied by a police detective. They recovered the computer and arrested two guys on the spot. They also found other "hot" items at the house. Sometimes you get lucky.
Blystone in Texas
-------------------------------- Robert V. Blystone, Ph.D. rblyston-at-trinity.edu
Department of Biology Trinity University 715 Stadium Drive San Antonio, Texas 78212 210.736-7243 FAX 210/736-7229
We would like to look at some heavier elements using the TEM, specifically alloys with Pt and Rh. Does anyone know the equation for calculating the thickness (thinness) requirements for samples to be electron transparent for different accelerating voltages?
TIA, John -- John Phelps NIST / DIV 853 Boulder, CO 80303
phelps-at-enh.nist.gov
ph. 303-497-5806 fax 303-497-5030
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