Dear microscopists, Does anybody know e-mail or fax of Agar Scientific in US? Thank you in advance.
___________________________ Dr. Alexander A. Mironov Jr. Unit of Morphology Dept. of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale, S.Maria Imbaro (Ch) 66030 Italy
Hello Alexander, I don't know the US details but the Agar Scientific head office in the UK will, their fax number is (+44) 1279 815106. Good luck,
Ron
On Sun, 1 Nov 1998, Alexander Mironov Jr. wrote: } } } Dear microscopists, } Does anybody know e-mail or fax of Agar Scientific in US? } Thank you in advance. } } ___________________________ } Dr. Alexander A. Mironov Jr. } Unit of Morphology } Dept. of Cell Biology and Oncology } Consorzio Mario Negri Sud } Via Nazionale, S.Maria Imbaro (Ch) } 66030 Italy } } Tel. 0872-570-332 } Fax 0872-578-240 } E-mail: amironov-at-cmns.mnegri.it } } } }
=========================================================================== 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. ============================================================================
Eastman Chemical Company has an opening for a Technician in the Physical Chemistry Research Laboratory. A candidate should have a 2-year degree, or higher, and experience in optical and/or electron microscopy. A background in chemistry is preferred but not required. The duties will include the use of optical and electron microscopes in support of research projects and in solving problems relating to products and manufacturing. The laboratory is fully state-of-the-art and offers varied and challenging assignments.
Eastman Chemical Company is one of the largest chemical, fiber and plastics manufacturing facilities in the U.S. The company's central research laboratories are in Kingsport, Tennessee, and provide a variety of resources for the technical community. Kingsport is located in northeastern Tennessee in the foothills of the Smoky Mountains. The area is rated among the top 25 most livable metropolitan areas and offers affordable housing, low taxes, and excellent schools. The pleasant four-season climate permits a variety of outdoor recreational opportunities.
Interested persons should send their resume to: Eastman Chemical Company, Employment Department, P.O. Box 1975, Kingsport, TN USA 37662-5215. Eastman Chemical Company is an Equal Opportunity Employer.
Dennis B. Barr (dennbarr-at-eastman.com) Physical Chemistry Research Laboratory Physical & Analytical Chemistry Research Division Eastman Chemical Company Kingsport, TN 37662-5150
In reference to embedding rubber from plants. So the rubber has been extracted from the plant and she wants to look at the suspension of rubber particles. Natural rubber or latex derived from plants is cis-polyisoprene -(CH2-C2H3=CH-CH2)- , which has an available double bond. If she has a suspension, I wouldn't extract, but just go ahead and dilute the latex, and then stain with a saturated bromine solution, or osmium tetroxide. A droplet can be placed on a carbon film on a grid. If cross-sections are necessary, cryo (as mentioned in an earlier E-mail) is an option. Or maybe she could dry a film, and harden the material with osmium prior to RT sectioning. Just thoughts on a Monday morning. Take care, Vicky
Does anybody know a supplier of the old-style scotch tape that can be used to make thin TEM samples? It can be used to remove thin layers of samples. You could easily make a nice graphite sample this way. I appreciate any help I can get. Thank you.
Peggy Bisher.
Margaret E. Bisher
NEC Research Institute 4 Independence Way Princeton, NJ 08540. Tel.: (609) 951-2629 Fax: (609) 951-2496 e-mail: peggy-at-research.nj.nec.com
Have you tried a different batch of protien a-gold? I've only done this kind of experiment once, a long time ago, but when I had problems this was the problem.
Karen Pawlowski
On Fri, 30 Oct 1998, Dwight Beebe 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 everyone, } I have encountered considerable difficulty, i.e., zero success, in } repeating an immunolabelling experiment. I'm working with LR White } embedded cotyledon tissue, fixed 12/95, probing with a polyclonal Ab } (rabbit), also made in '95. My secondary is commercial protein A gold. I } had very specific labelling when the initial experiment was conducted in } '95, but now have no specific label. I have repeated the previous } successful protocol, but with no luck. My antibodies have been at -20C, } having only been thawed once to aliquot them into smaller volumes. } Is it possible that there have been changes in the fixed tissue } that cause loss of epitopes? I find this hard to believe, but I'd like to } hear from others. Could the antibodies have lost their specificity? I } have repeated dilution series experiments, but again, all labellings were } no more specific than buffer controls. I should say that the antibodies } work fine in Westerns, giving us the same banding pattern we've always seen } with these Ab, so I suspect the tissue, not the Ab. } Any and all comments, criticisms, speculative remarks, SWAGs, etc. } would be most welcome. Much hinges on the success of the labelling. } Many thanks, } Dwight } } } ************************************************************************* } Dwight Beebe } Prof. Agrege (Associate Prof.) } Institut de recherche en biologie vegetale } Universite de Montreal } 4101 est, rue Sherbrooke } Montreal, (Quebec) H1X 2B2 Canada } Tel: 514/872-4563 or -4746 (lab) } FAX: 514/872-9406 } } } }
When this kind of thing happens to me, I recheck the antibody on fresh frozen or what ever Light Level tissue sections you know it should work on. If the staining pattern is the same as the original experiment, then it would lead you to the tissue. I have seen LRWhite tissue sections become unusable in matter of days or weeks sometimes, but the blocks have been pretty stable. I have also seen some strange staining pattern when the pH of the buffer is off.
Good luck!
Derm Imaging Center University of Washington
On Fri, 30 Oct 1998, Dwight Beebe 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 everyone, } I have encountered considerable difficulty, i.e., zero success, in } repeating an immunolabelling experiment. I'm working with LR White } embedded cotyledon tissue, fixed 12/95, probing with a polyclonal Ab } (rabbit), also made in '95. My secondary is commercial protein A gold. I } had very specific labelling when the initial experiment was conducted in } '95, but now have no specific label. I have repeated the previous } successful protocol, but with no luck. My antibodies have been at -20C, } having only been thawed once to aliquot them into smaller volumes. } Is it possible that there have been changes in the fixed tissue } that cause loss of epitopes? I find this hard to believe, but I'd like to } hear from others. Could the antibodies have lost their specificity? I } have repeated dilution series experiments, but again, all labellings were } no more specific than buffer controls. I should say that the antibodies } work fine in Westerns, giving us the same banding pattern we've always seen } with these Ab, so I suspect the tissue, not the Ab. } Any and all comments, criticisms, speculative remarks, SWAGs, etc. } would be most welcome. Much hinges on the success of the labelling. } Many thanks, } Dwight } } } ************************************************************************* } Dwight Beebe } Prof. Agrege (Associate Prof.) } Institut de recherche en biologie vegetale } Universite de Montreal } 4101 est, rue Sherbrooke } Montreal, (Quebec) H1X 2B2 Canada } Tel: 514/872-4563 or -4746 (lab) } FAX: 514/872-9406 } } } }
Interface roughness in GaAs/AlGaAs was a hotly debated subject a few years back. Optical techniques seemed to indicate a perfect, atomically flat interface with occasional steps, HRTEM showed roughness at the atomic scale. I was involved with the HRTEM part. We used a technique developed by Ourmazd et. al. at Bell Labs to analyze the HRTEM cross sections. We called it "chemical mapping" or "chemical lattice imaging". I don't know if this helps, as you don't say, what exactly your interface consists of, but try a literature search for the above keywords. Also try to search for "Ourmazd". Other authors on various publicatons were Kim, Baumann, Warwick, and myself. I have been out of the field for a few years, but Ourmazd and coworkers have extended the analysis to SiGe systems as well.
The software we used was developed by a number of people at Bell Labs and ran on a Silicon Graphics. At the time we were giving away the software to other researchers. I don't know, if that is still the case. The group at Bell Labs does not exist anymore, Ourmazd is now in Frankfurt/Oder in Germany.
Good luck.
Michael Bode Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 phone: (888) FIND SIS fax: (303) 234-9271 email: info-at-soft-imaging.com
} } } ---------- } From: Philip Flaitz[SMTP:flaitz-at-us.ibm.com] } Sent: Friday, October 30, 1998 1:05 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: TEM: interface roughness analysis. } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
This is a summary of the replies I rec'd to my inquiry about beam damage to carbon tape. Thanks to all who replied.
Owen
+++++++
My initial message was:
Good morning:
We jet spray volcanic ash particles (~10 microns or less) onto carbon double stick tabs for SEM/EDS study. It is easy to produce a nice distribution of particles with few that overlay one another. Problem is that later, during EDS and backscatter imaging, when the beam current is higher, the carbon tape is often damaged. It curls and bubbles thereby distorting the images.
We tried to use carbon paint but it is tricky to get the right amount spread onto the mount or it dries too quickly. Results were unfavorable.
I'd really like to have a durable, very smooth, low Z surface, that is insensitive to beam current and is easy to apply and work with in the SEM. Oh, it should also be economical too.
Any ideas? TIA
Owen
+++++++++
The carbon tape is successful with distributing the effects of charging but not heat. I, at least, can only suggest you sputter with a heat conducting metal ... e.g., Al, Au, Cu ... most any metal would help, but I realize it would superimpose spectra and degrade your BSE contrast. Short of that, you'd need to lower the energy in your beam with either lower beam currents or accel voltage. I'll also be looking for other ideas ... hope this helps :o)
cheerios, shAf 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/
+++++++++
I have never had damage to carbon tape, even at 30kV. I store mine in the fridge when not in use, as I heard it deteriorates. I bought my rolls from SPI several years ago and never had a problem.
Alan Stone ASTON Metallurgical Services
+++++++++
I would like to suggest Berylium as a possible solution for a smooth, low-Z surface. It can be evaporated onto the specimen and it satisfies all the given criteria. HOWEVER, there are very serious safety issues involved. If you have access to the appropriate equipment--a dedicated vacuum evapora- tor with exhaust vented appropriately, the equipment to produce small Be pellets or wire for evaporation, a safe place to clean out the bell jar, etc.--you might want to consider using Be. I think that solid Be metal is not too dangerous, and that finely-divided metal or compounds are the most hazardous. I have handled BeO powder (using a hood, of course) and had no trouble, and I heard a presentation at MSA some years ago by Dr. Hall (whose first name escapes me) where Be coating was used instead of C for low temper- ature work (where Be's conductivity is much greater than C's). With the stated safety reservations, perhaps you could produce a Be equivalent of double-stick tabs. Good luck. Yours, Bill Tivol
++++++
Owen,
Have you tried double sided conductive carbon sheets, instead of the tabs? While the systems appear to be similar, the sheet product is a bit more resistant to changing in the electron beam. Now if that either fails as well, or is not something you would want to try, there is a method I have described on the listserver at least once before and maybe twice. It is based on the use of a camphor naphthene eutectic composition. For the coverglass smoothnes, purchase HOPG materials. would seem like for your appliction, being non-SPI, that the cheapest grade would be acceptable for you. Remember that is just some judgement on my part but it might not be correct. But you want to make a freshly cleaved surface to expose the nice smooth surface of HOPG. I beam won't hurt it in the least. I guarantee that. Then use the camphor naphthalene procedure to disperse the particles on the HOPG. A quick and dirty verion of this might be to put the HOPG on a hot plate at something around 100 deg. C, and disperse your ash particles in acetone, the acetone will evaporate away before the particles have time to agglomerate. But if they don't, then you should turn to the camphor naphthalene method.
Chuck
++++++++
Ihave quite often used conductive carbon tape for such exams with no problem.
.Wonder what beam current and voltage you are running? I presume you are carbon coating the ash/tape to aviod charging and that is not the distortion
problem.
Woody White McDermott Technology, Inc. +++++++
Owen, Since you don't need much adhesive to tack down 10 micron particles, you may try applying a very thin layer of a fluid adhesive (such as Microstick) to a pyrolytic carbon planchet. This form of carbon has a flat, hard, glass-like surface. A drop of adhesive applied to the carbon may be spread very thin by drawing out with a cover slip. Since the organic adhesive layer is thin, you will probably get your required conductivity. There is no observable background structure. Planchets are about $35 each.
Dennis.
Dennis C. Ward voice: 202-324-2982 FBI fax: 202-324-4018 Microanalysis Laboratory e-mail: DCWard-at-concentric.net
+++++++++
The only way to do this is with Carbon Dag , what you do is take a match dip it into the dag and with the match level to the stub just pull across stub only pull match across once this ensure quite a flat level. Just be sure to dilute the dag quite a bit this gives you more of a flat base.
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
++++++++
As far as the tape goes, we use a 3M double back tape that seems to work fairly well so long as you carbon coat the samples. You may want to give it a try. We have also developed an ethyl acetate copolymer that we apply to mylar film which seems to work even better.
Keith Rickabaugh Manager, Materials and Particle Characterization {krickabaugh-at-rjlg.com}
RJ Lee Group, Inc. 350 Hochberg Road Pittsburgh, PA 15146 ph: 724-325-1776 {www.rjlg.com}
+++++++++
Dennis has a good idea. I would substitute freshly-cleaved mica for the carbon planchet...its a lot cheaper.
Chuck Butterick Engineered Carbons, Inc .
+++++++++
There is a product that might fill your bill. It is a thermoplastic called Tempfix marketed by Electron Microscopy Sciences (EMS). In '94 we published a paper in Trans. Am. Microsc. Soc."Mounting Pollen on a Thermoplastic Adhesive for Scanning Electron Microscopy" in which we describe our protocol and demonstrate its advantages for our use. While we did not do any x-ray or BSE imaging on those samples, I believe it would hold up to such use (depending ofcourse on the accel. voltage and beam current used). If you cannot find the journal, I would be glad to send you a reprint if you're interested. Please contact me off line. I have no interest in EMS except as a customer.
Bill
+++++++++
Remember also that if you are not concerned about EDS peaks from the substrate background, consider Tacky Dot Slides. Those are really the ideal kind of mounting system if you are trying to catalog what you have in some quantitative basis, and they also make it easy to come back to the same particle characterized previously.
Chuck
+++++++
Owen
There IS an adhesive that is: smooth-surfaced, high tack, low Z, durable, easy to use, dependable, nontoxic and inexpensive. It is also pretty stable under the beam and thin layers do not outgas noticeably in the vacuum. It is, however, not conductive (after all - is there such a thing as a perfect whatever?). We use this stuff regularly to stick down pollen, fly-ash, small insects, blood cells, ceramic powders, pigment particles etc. for imaging and for EDS analysis.
it is called 'Artists Gold Size' and is a thick tacky syrup of partially polymerised liseed oil. On exposure to air it polymerizes and forms an insoluble polymer (this was used in the manufacture of old-style floor linoleum, hence the name linoleum - from 'linseed oleum').
Buy this at any art supply shop, spread a very small drop of the syrup over the surface of a stub, wait until it is just tacky enough not to wick up the sample, drop your volcanic ash particles onto the surface and leave for 30 min. at room temp (or a few min at 50C) to polymerize, coat with carbon and analyse.
I've never been able to find out who used this first, but have been using it for many years after coming across a casual reference to its use as a SEM mountant.
A $5 bottle should last you years.
Pity it isn't conductive though.
Jan C
============================= Owen P. Mills Michigan Technological University Metallurgical & Materials Engineering Rm 512 MME Building Houghton, MI 49931 906-487-2002 906-487-2934 FAX opmills-at-mtu.edu
Tannic acid is usually used with both the primary and the post fixation in 1-8% (w/v) concentration. The protocol I used successfully before was as the follows.
The fresh made solution of 12% tannic acid (Polysciences cat#4459, EM grade) buffered in 0.2 M cacodylate, pH 7.4 was mixed in an equal volume ratio immediately before fixation with 5% glutaraldehyde in 0.2 M cacodylate buffer, pH 7.4. The final pH is adujsted to 7.2 with 10 N NaOH. The tissues are then postfixed with 2% OsO4 only in the same buffer or a mixture of tannic acid with OsO4.
Good luck,
Ming
} Has anyone ever used tannic acid in their gluteraldehyde? I've heard it's } suppose to help preserve cilia and fibers. Any receipes, methods, or } concentrations would be appreciated. Thanks in advance.
*********************************************** * Ming H. Chen, PhD * * Medicine/Dentistry Electron Microscopy Unit * * #1074B Dentistry Pharmacy Building * * University Of Alberta. * * Edmonton, Alberta, Canada T6G 2N8 * * * * Visit My Page At: * * http://www.ualberta.ca/~mingchen * ***********************************************
This is one of those challenging imaging problems. There are several companies, including Wyko (now Veeco) and Zygo, which specialize in surface structure analysis. Both do 3-D interferometry. I am not sure that they are able to see throughh one layer into another and measure the roughness of the second surface, but it is worth a try. Both have web sites. If you can't get info, contact me off-line and I will supply contact names and numbers.
Dear Mike, When you do measurements in Image Tool, you should just hold down the mouse at the first point and release it at the second. The double-click is confusing the software. I am using the latest version (1.28) on the site. You wrote: } } I am using Image Tool software. When I do measurements by drawing the } marker and double clicking to end the measurement the program sometimes } crashes. Has this happened to anyone else? Is so how do you correct } the problem? } Michael Ingram } Rodel, Inc
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
Hi I have neural tissue embedded in Unicryl by way of UVlight at 4 degrees C. It was fixed with 4% paraform and 0.5% glut, no osmium. I am having difficulty staining with 2% aquas Ua and Renolds lead citrate. I have tried combinations from 3 min to as much as 10 min with very little luck, yet the same stains work with my standard 812 embedded sections at 6 min and 5 min respectively. My past experience with LRW was that it took less time and or concentrations than epon like plastics. The T-Blue stained very quickly like LRW. 1. Anyone out there have some experience or ideas? 2. What kind of time periods should one use for washes in drops of distilled water after Ua?
We would like to attach a small hydrophobic protein to small gold particles (less than 5nm). The problem is that most recipes are for proteins in aqueous solutions. We would like to put the gold into chloroform/methanol since our protein is happiest in that, and have the labelling proceed. Does anyone have experience with this?
We have tried mixing protein and gold each in 70% isopropanol since both are happy in that; but we get precipitate when we dry the mixture down.
Gold can be prepared (and obtained from Nanoprobes) with attached alkanes to which our protein might be adsorbed but we would like to avoid increasing the size of the particles by this intermediate layer and further wonder if such an extra layer might not weaken the protein-gold linkage, the protein come off when added to the tissue and gold stick non-specifically to other things.
We can't do immunogold since the protein is likely buried in lipid bilayers and has not been possible to label with conventional aqueous antibody-gold. The protein is surfactant protein B (Possmayer, Voorhout, Hawgood).
Since the gold-protein bond is said to be hydrophobic (and the degree of hydrophobicity of a protein to correlate with its stickability) we hope that the gold should stick to the protein.
Any thoughts will be appreciated, Jacob
Jacob Bastacky, M.D. Room 116 Donner Laboratory Lawrence Berkeley Laboratory University of California Berkeley, California 94720 Telephones: 510.486.4606 office, 510.486.4605 lab, 510.845.8031 fax email: sjbastacky-at-lbl.gov
6 weeks ago I posted an inquiry about color slide photography of computer screens. Lots of helpful replies made my little project an easy success.
I used Kodak EliteChrome ASA 100, f5.6 to 8, 0.5 second exposure and midrange brightness and contrast on the monitor. A tripod and cable release were employed. The camera was placed about 3 feet from the screen and a 58mm lens was used.
The resulting slides showed faithful color and were sharp enough to pass for original slides to the unsuspecting.
The main difficulties were in framing, and in distortions relating centering and perpendicularilty of the camera.
A black mat cut to the aspect ratio of the camera was used to solve the framing problem.
The distortions were more troublesome and not entirely overcome. At three feet slight off axis camera placement creates noticeable taper. A carefully made perpendicular "hood" mount would help. Bill Miller suggested mounting a small mirror on the center of the screen and looking for a reflection right back down the lens as a means of orienting the camera. I didn't try this. Moving farther away and using a telephoto lens is another suggestion which I didn't try, but should have.
Email: TKGreen-at-aol.com Name: Tristan Green School: Piedmont High School
Question: Do you know of a good place to find pictures of common bacterial cultures to use for identification purposes? We are culturing colonies on petri dishes from swabs taken around campus. I would love for my students to be able to identify what they've found.
Thank you, Mrs. Tristan Green Science Teacher, Piedmont High School
6 weeks ago I posted an inquiry about color slide photography of computer screens. Lots of helpful replies made my little project an easy success.
I used Kodak EliteChrome ASA 100, f5.6 to 8, 0.5 second exposure and midrange brightness and contrast on the monitor. A tripod and cable release were employed. The camera was placed about 3 feet from the screen and a 58mm lens was used.
The resulting slides showed faithful color and were sharp enough to pass for original slides to the unsuspecting.
The main difficulties were in framing, and in distortions relating centering and perpendicularilty of the camera.
A black mat cut to the aspect ratio of the camera was used to solve the framing problem.
The distortions were more troublesome and not entirely overcome. At three feet slight off axis camera placement creates noticeable taper. A carefully made perpendicular "hood" mount would help. Bill Miller suggested mounting a small mirror on the center of the screen and looking for a reflection right back down the lens as a means of orienting the camera. I didn't try this. Moving farther away and using a telephoto lens is another suggestion which I didn't try, but should have.
The TEM group of Abbas Ourmazd has moved to the Institute for Semiconductor Physics in Frankfurt (Oder), Germany. The group continues to apply quantitative HRTEM and electron holography. Peter Schwander is in charge for maintaining the Chemical Mapping and QUANTITEM software. This software is ready to run on SGI workstations under IRIX 5.3 and 6.2.
Anyone interested in Chemical Mapping or QUANTITEM or any related information is welcome to contact Peter Schwander ( schwander-at-ihp-ffo.de ).
Alex *=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*= =B0* Dr. Alexander Orchowski Institute for Semiconductor Physics Frankfurt (Oder), Germany http://www.ihp-ffo.de phone: +49 (0) 335 562 5432 fax: +49 (0) 335 562 5300 email: orchowski-at-ihp-ffo.de *=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*=B0*= =B0*
Michael Bode wrote:
} Phil,
} Interface roughness in GaAs/AlGaAs was a hotly debated subject a few } years back. Optical techniques seemed to indicate a perfect, atomically } flat interface with occasional steps, HRTEM showed roughness at the } atomic scale. I was involved with the HRTEM part. We used a technique } developed by Ourmazd et. al. at Bell Labs to analyze the HRTEM cross } sections. We called it "chemical mapping" or "chemical lattice imaging". } I don't know if this helps, as you don't say, what exactly your } interface consists of, but try a literature search for the above } keywords. Also try to search for "Ourmazd". Other authors on various } publicatons were Kim, Baumann, Warwick, and myself. I have been out of } the field for a few years, but Ourmazd and coworkers have extended the } analysis to SiGe systems as well.
} The software we used was developed by a number of people at Bell Labs } and ran on a Silicon Graphics. At the time we were giving away the } software to other researchers. I don't know, if that is still the case. } The group at Bell Labs does not exist anymore, Ourmazd is now in } Frankfurt/Oder in Germany.=20
} Good luck.
} Michael Bode } Soft Imaging System Corp. } 1675 Carr St., #105N } Lakewood, CO 80215 } phone: (888) FIND SIS } fax: (303) 234-9271 } email: info-at-soft-imaging.com =20 } Philip L. Flaitz wrote:
} } I have had a request to examine an interface structure which, while nominally } } planar, has roughness associated with it. In addition to imaging the } } structure, the person making the request would like to have a measurement= of } } the roughness associated with the interface. The interface of interest is } } etched within Si and thus inaccessible to AFM.
} } Is anybody aware of techniques for measuring interface roughness from a= TEM } } image? Are there any software packages which have such function built-in?
} } Phil
} } Philip L. Flaitz } } IBM Analytical Services } } Ph.......(914) 892-3094, FAX -2003 } } flaitz-at-us.ibm.com
Something we and others have experimented with is the use of Dimethylsulfoxide (DMSO). It acts as a cell penetrant and stabilizes the cytoskeleton. My work is principally with human neutrophils but others have used it for yeast cells (and other things, but this should be enough to test the idea.)
References:
Gilbert, CS and RT Parmley Morphology of human neutrophils: A comparison of cyrofixation, routine glutaraldehyde fixation, and the effects of dimethyl sulfoxide Anat Rec 252:254-263 (1998)
Fassel, TA; Sohnle, PG and VM Kushnaryov The use of dimethylsulfoxide for fixation of yeasts for electron microscopy Biotechnic & Histochemistry 72(5):268-272 (1997)
Forgive my Portuguese, it's little rusty. I can help with acquiring copies of the references if you need them.
Ciao,
Chuck
--- On Wed, 28 Oct 1998 20:37:25 -0200 Rinaldo Pires dos Santos {rinaldop-at-botanica.ufrgs.br} wrote: ------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Dear colleagues
I am working with the ultrastruture of anthers of Ilex paraguariensis. However, during the dehydration of the samples, fixed in a mixture of glutaraldehyde 2,5% and formaldehyde 2%, the anthers shrink, mainly after the secondary fixation with osmium tetroxide. I used acetone or ethanol , in steps of 30, 50, 70, 90, 90, 100, 100, with 15 minutes in each step, at room temperature. The anthers has a very reduced dimension (about 1 mm length) and shrink in the step 100. What to do? Should I to use a low temperature (4oC)? Thanks in advance.
-----------------End of Original Message-----------------
------------------------------------- Name: Charles Gilbert VOC:(704)355-5261 Carolinas Medical Center FAX:(704)355-8424 Dept of Pediatric Research PO Box 32861 Charlotte, NC 28232-2861
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
On Fri, 30 Oct 1998, Alexander Mironov 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 } -----------------------------------------------------------------------. } =20 } =20 } Dear microscopists, } We are planning to buy some diamond knives and I would like to know your } impressions of knife performance (what is the difference between Diatome, } Drukker, Pelco, DDK etc.). We need standard ultramicrotomy knives for } cutting epon embedded biological material and cryo dry ultramicrotomy } knives. Personally, I like Diatome and dislike DDK. } Did anybody see the difference in cutting and durability between 45=B0 an= d } 35=B0 knives?=20 } Which boat for standard knife is preferable? What is the purpose of boats } with inclined cavity? } Any experience is wellcome. } =20 } ___________________________ } Dr. Alexander A. Mironov Jr. } Unit of Morphology } Dept. of Cell Biology and Oncology } Consorzio Mario Negri Sud } Via Nazionale, S.Maria Imbaro (Ch) } 66030 Italy } =20 } Tel. 0872-570-332 } Fax 0872-578-240 } E-mail: amironov-at-cmns.mnegri.it } =20 } =20 } =20 } =20 I have used DDK, Dupont, Diatome, others. For the last ten years we have invested only in Diatome. For the last seven years we have about 22 thousand dollars worth of assorted Diatome knives. We have Never had a bad one, never had one poorly resharpened (I long ago quit testing them when they came back to us), never had one that wore out quickly. Furthermore, Diatome USA has a laboratory set up. In case you have trouble with your embedding or materials, you can send them some of your tissues with your knife, and they will probe the situation for you and give you advice. I have not used this service, but I know from others that this service is fine. We have gotten excellent advice also on solving problems with our microtomes which was affecting the knives. I don't own stock in Diatome. Wish I did.
Since the release of Time magazine's Oct. 12th article concerning the Deep Space I project, which is now heading for an asteriod orbiting the sun, we have been inundated with inquiries concerning our electron microscope apertures. Let me try to answer some of the questions posed to us. The xenon ion propulsion system (XIPS) has launched a new era in satellite propulsion technology. Currently three satellites are in geo-synchonous orbit above the earth using XIPS. In conjunction with the satellite designers we have been able to adapt the standard electron microscope aperure in the thruster control devices in the XIPS, which Time magazine described as the " stuff of technological fantasies ". In addition to EM aperures with non-standard holes, ultrathin and special metals, EN type apertures are also now being used in other applications, including:
- FIBS - X-ray collimators and focusing devices - Light and gas control devices requiring precise microholes and slits - Solder droplet production
We believe the advances we have made in these other applications have allowed us to improve EM apertures overall. I hope this answers most of your questions. We are sending out more detailed information to those who requested it.
Thank you for your interest,
John Arnott Chairman --
LADD RESEARCH 13 Dorset Lane Williston, VT 05495
TEL 1-800-451-3406 (US) or 1-802-878-6711 (anywhere) FAX 1-802-878-8074 e-mail ladres-at-worldnet.att.net web site http://www.msa.microscopy.com/SM/LADD
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On Fri, 30 Oct 1998, Alexander Mironov 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 } -----------------------------------------------------------------------. } =20 } =20 } Dear microscopists, } We are planning to buy some diamond knives and I would like to know your } impressions of knife performance (what is the difference between Diatome, } Drukker, Pelco, DDK etc.). We need standard ultramicrotomy knives for } cutting epon embedded biological material and cryo dry ultramicrotomy } knives. Personally, I like Diatome and dislike DDK. } Did anybody see the difference in cutting and durability between 45=B0 an= d } 35=B0 knives?=20 } Which boat for standard knife is preferable? What is the purpose of boats } with inclined cavity? } Any experience is wellcome. } =20 } ___________________________ } Dr. Alexander A. Mironov Jr. } Unit of Morphology } Dept. of Cell Biology and Oncology } Consorzio Mario Negri Sud } Via Nazionale, S.Maria Imbaro (Ch) } 66030 Italy } =20 } Tel. 0872-570-332 } Fax 0872-578-240 } E-mail: amironov-at-cmns.mnegri.it } =20 } =20 } =20 } =20 I have used DDK, Dupont, Diatome, others. For the last ten years we have invested only in Diatome. For the last seven years we have about 22 thousand dollars worth of assorted Diatome knives. We have Never had a bad one, never had one poorly resharpened (I long ago quit testing them when they came back to us), never had one that wore out quickly. Furthermore, Diatome USA has a laboratory set up. In case you have trouble with your embedding or materials, you can send them some of your tissues with your knife, and they will probe the situation for you and give you advice. I have not used this service, but I know from others that this service is fine. We have gotten excellent advice also on solving problems with our microtomes which was affecting the knives. I don't own stock in Diatome. Wish I did.
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Dear Mr. Corwin,
1) Encapsulate the sample fully in Epoxy 2) Using a high speed diamond saw, cut the sample with a RESIN Bonded diamond blade to minimize chipping of the edge. 3) If the cut surface is not good enough, a quick polish maybe necessary.
Try HiRel Laboratory in Spokane, Washington, or Metals Technology in Northridge, Ca. Either of them can be found in directory assistance.
Good Luck,
Gary Liechty Allied High Tech Products, Inc. Products for Metallographic, SEM and TEM Sample Preparation 800-675-1118
} ------------------------------------------------------------------------ } 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 suggest how to cut into a glass pharmaceutical vial with } minimal damage and contamination in order to examine an organic film } inside it? Presumably some sort of diamond saw. I need a commercial } lab who can provide the service, ASAP of course. Hatchet is last } resort. Thanks for your help. } } } Leonard Corwin } Research Chemist } Fort Dodge Animal Health } Princeton, NJ 08543-0400
--------------87761FD3C5399E2310A98724 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"
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I am attempting to help a colleague who is in a bind. She needs to get QUICKLY - which is to say - borrow, loan, rent, or buy - a computer controlled stage for an Olympus BH-2 optical microscope.
We're talking to Olympus of course but still ...
Anyone have ideas or a stage available???
Richard Shalvoy rbshalvoy-at-corp.olin.com 203-271-4272
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Hey There Listers,
I have a user who wants to embed rubber extracted from plants. The last time she did it she used glut & OsO4 (concentrations I do not know). The insides of the what she thought were solid rubber particles were extracted or at least they looked extracted (so the samples kinda looked like an inner tube). Do any of you bouncing baby scientists have any suggestions for her? She says this stuff is lipid-like and is a short-chain rubber compound, so she figures its pretty liquid.
Send me your suggestions & I'll bounce them off her & see if any stick.
Paula :-)
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
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We are interested in using the SEM to measure the concentration of flyash in the atmosphere that has come from coal-fired power plants. One of the issues involved is being able to distinguish particles from power plants from particles originating at other sources. We would appreciate any information people might have on the identification of flyash and determination of its origin. Everett Ramer Federal Energy Technology Center
Has anyone ever tried any Butvar other than 98? I found a jar of Butvar-79 (from Monsanto) in our chemical cabinet and was thinking of giving it a whirl. I'm guessing it is a shorter polymer than the 98. Nobody knows what it is doing here...it seems to have been inherited from the previous inhabitants.
By information about SEM in seeds of Avena sativa and Avena byzantina (oats). Thanks very much
Diana Reinoso
cultivar-at-fca.uner.edu.ar Facultad de Ciencias Agropecuarias Universidad Nacional de Entre R=EDos Rep=FAblica Argentina --------------------------------- Catedra Botanica Sistematica Fac. Cs. Agropecuarias UNER C.C. 24 3100 - Parana - Entre Rios ARGENTINA Tel: 043 - 975075 Fax: 043 - 975096 E-MAIL: botanica-at-fca.uner.edu.ar
Here I am once again, hat in hand to ask for your help. This time I have someone who wants to do SEM on a cell line grown in suspension (they are completely non-adherent, so we can't grow them that way). What should we do in terms of fixation, etc. and then how do we stick them to the stub? Do we try to make them stick to something first & then run them up? Do you run them up first & then stick them to something? Do you run up to them & say "Stick 'em up"? As you can tell, this is a new one to me. I've process little things by wrapping them in lens tissue & then running the up & then shaking the sample onto the sticky carbon dots. I don't think that procedure will work for these because these cells are much tinier. I thank you in advance for any suggestions you can give me. And I thank you for all the help and advice you've given me in the past.
I'll stick by my computer until I hear from y'all,
Paula :-)
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
1 a supplier of cheap glass slides about 26 x 46 mm, 1 to 1.5mm thick?
2 someone who can take a given powder/dust sample and perform a respirable/non-respirable binary split of it and return to me the two size fractions for subsequent characterisation?
thanks
Ritchie
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
Postdoctoral Position in Interfacial Segregation Department of Chemical and Materials Engineering University of Kentucky Lexington, KY USA
A postdoctoral position is available in the area of interfacial segregation. The research project is focused on understanding interfacial segregation at ceramic/ceramic and metal/ceramic interfaces and it effects on physical properties of interfaces. Through a collaboration with Prof. Susan Sinnott, a computational materials scientist at the University of Kentucky, the issue of interfacial segregation will be addressed from a combined experimental and theoretical approach. The ideal candidate for this position will have experience in HREM, STEM , EDS and/or EELS. The University of Kentucky is in the process of developing a state-of-the-art microscopy facility and will be purchasing a 200kV Field Emission TEM/STEM in 1999. The University has an active and growing materials science program with many interdiscplinary research and educational activities between the Materials Engineering, Physics and Chemistry Departments and the Center for Applied Energy Research. Many opportunities for performing experiments at Oak Ridge National Laboratory are also available. Please send applications to Professor Elizabeth Dickey at the address below.
***************** Professor Elizabeth Dickey Department of Chemical and Materials Engineering 177 Anderson Hall University of Kentucky Lexington, KY 40506-0046 USA ph: 606.257.8572 FAX: 606.323.1929 email: ecdickey-at-engr.uky.edu
We can't leave you dangling like this. There are several approaches to solve this suspensful situation. I base this on work I have done with bacteria, fungi and tissue cultures grown in suspension.
In the first approach (easy but yielding few, isolated cells) , take a scrupulously clean, glass microscope slide (use a good glassware detergent and rinse it quite well - do not use acid cleaning agents as they may be toxic). Treat the glide with a poly-L-lysine solution (1 mg/ml distilled water) for 5 minutes, then rinse with distilled water. Place slide into a petri dish containing a filter paper moistened with distilled water and containing several pieces of applicator stick. The slide should be suspended (like a bridge) over the applicator sticks. Gently layer on the cell suspension on the slide so that it fills the slide but does not overflow the edges (about 5 ml). Allow the cells to settle onto the slide for about 1 hr. You should keep the cells "happy" during this time period (out of the light, warm, perhaps in a CO2 incubator). Now, very carefully aspirate away most of the liquid and very slowly add 5 ml of your primary fixative (4% formaldehyde/2%glutaraldehyde) to again cover the slide. Allow to fix for 15-20 min at room temp. Rinse by gently decanting and gently adding buffer from one end of the slide. Post-fix in 2% osmium tetroxide (buffered or in distilled water) for 30 min at room temp. Rinse in distilled water and dehydrate the entire slide by overlaying with 50, 75, 95, ABS ethanol. They key here is "gently". After absolute ethanol, critical point dry the slide (or portion of the slide), coat with conductive metal and examine in the SEM. This technique will give you beautiful black backgrounds with only a few cells per field of view. Some people prefer this method since the cells are quite isolated and not piled up.
A second approach (possibly yielding too many cells on a substrate), is to pass the cell suspension through a micropore filter or silver membrane (available through Structure Probe, for example) so that the cells are retained on the surface of the membrane. This must be done very gently so that the cells are not broken. We have best success by sucking the cells down onto the membrane rather than forcing them onto the membrane from a syringe. You will need to try several dilutions of cell suspension since too many cells could pile up and create a mess. After filtering the cells, you would then apply the various solutions (fix, wash, dehydrate) by passing through the filter. In the end, you would remove the filter and transfer it into the CPD device (keeping it wet, of course). Some holder will be needed to help keep the filters from flip flopping in the CPD device and to maintain the proper orientation (i.e., keep track of which side the cells are on). Mount the filter on the stub (double sticky tabs), coat with metal and examine in the SEM.
Good luck,
JB
} Here I am once again, hat in hand to ask for your help. This time } I have someone who wants to do SEM on a cell line grown in suspension (they } are completely non-adherent, so we can't grow them that way). What should } we do in terms of fixation, etc. and then how do we stick them to the stub? } Do we try to make them stick to something first & then run them up? Do } you run them up first & then stick them to something? Do you run up to } them & say "Stick 'em up"? } As you can tell, this is a new one to me. I've process little } things by wrapping them in lens tissue & then running the up & then shaking } the sample onto the sticky carbon dots. I don't think that procedure will } work for these because these cells are much tinier. } I thank you in advance for any suggestions you can give me. And I } thank you for all the help and advice you've given me in the past.
#################################################################### 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 ####################################################################
Does anyone have a reference for nuclear microclefts in the journal "Ultrastructural Pathology"? I'm sure I read about them in this journal three or four years ago but cannot find the reference now despite a Medline search and a look through our journals. Nuclear microclefts are sometimes found in small cell carcinomas. I am trying to establish how specific this observation is in the setting of tumour diagnosis.
Thankyou.
John Brealey, EM Unit, The Queen Elizabeth Hospital, Adelaide, South Australia.
Paula Sicurello wrote: =============================================== Here I am once again, hat in hand to ask for your help. This time I have someone who wants to do SEM on a cell line grown in suspension (they are completely non-adherent, so we can't grow them that way). What should we do in terms of fixation, etc. and then how do we stick them to the stub? Do we try to make them stick to something first & then run them up? Do you run them up first & then stick them to something? Do you run up to them & say "Stick 'em up"? As you can tell, this is a new one to me. I've process little things by wrapping them in lens tissue & then running the up & then shaking the sample onto the sticky carbon dots. I don't think that procedure will work for these because these cells are much tinier. ============================================== We have had over the years heard good reports from people who have filtered onto silver membrane filters (they cost about 3X that of a normal polymer membrane filter), the cells are kept wet, and then processed for critical point drying. They are then CPD'd and looked at by SEM. Because of the highly conductive nature of the substrate, much less gold, if any is then required, an advantage for particularly fine structured or heat sensitive samples.
A second approach, but one that I am not as enthusiastic about, but one which will work just the same, is the use of microporous specimen capsules , which have a pore size that is as small as 30 um. You can CPD the cells right in the capsules, and then when dry, with a razor blade, you can slit open the capsules, put on your conductive coating, and examine the cells in situ, right on the surface of the now slit open microporous specimen capsule
Dear Everet } } We are interested in using the SEM to measure the concentration of } flyash in the atmosphere that has come from coal-fired power plants. } One of the issues involved is being able to distinguish particles from } power plants from particles originating at other sources. We would } appreciate any information people might have on the identification of } flyash and determination of its origin. } Everett Ramer } Federal Energy Technology Center
I did a lot of SEM on Flyash some years ago which originated from different power plants. Thy were always perfect spheres. I kept some for atigmatism correction test samples. By doing EDS we were able to trace them back to the original power plants by comparing trace elements which were present in the Flyash 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
I have just heard of an elderly TEM in full working condition which is available free to anyone who is prepared to take it away. It is a Siemens Elmiskop 102 complete with Siemens image intensifier, and has had both HT cables replaced relatively recently. The instrument is installed in the north of England.
If anyone is interested, please contact me directly.
My only commercial interest is that as an ex-Siemens TEM engineer, I would be pleased to assist in the dismantling etc. if required.
Paula, Cells will adhere to coverslips or silica chips treated with poly-L Lysine (Sigma Aldrich) or suspended cells can be caught using a 0.2um polycarbonate filter. Cells should be fixed following either an incubation on the treated substrate to allow for attachment or immediately after filtering. I open the filter holder and transfer the filter to a specimen processing holder filled with fixative (EMS - cat.# 70186) Poretics (800-922-6090 ) is a good source for the 13mm Swinney syringe holders and the track etch filters. They also have a helpful specialist in Charlotte Hargrave. Rosemary
a German company called Maerzhaeuser sells motorized stages for the Olympus BH-2. Models 100 through 150 fit this Microscope. If you need more information, please contact me at my email address below.
Michael Bode
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 phone: (888) FIND SIS fax: (303) 234-9271 email: info-at-soft-imaging.com
************************************************************************ ********************************** Disclaimer: As we sell image processing systems including this stage, I do have a commercial interest in this. ************************************************************************ ******************************************************* } } ---------- } From: Shalvoy, Richard[SMTP:rbshalvoy-at-corp.olin.com] } Sent: Thursday, October 29, 1998 2:36 PM } To: Microscopy-at-sparc5.microscopy.com } Subject: Need computer controlled microscope stage } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I am planning to inject an antibody conjugated with ultrasmall gold particles into a living mouse, fix by cardiac perfusion, and embed the tissues for examination by both LM and TEM. The gold particles will be silver enhanced on the sections as even our EM studies are done at rather low magnification. In the past I have done this with antibodies conjugated to 125-I, then fixed with glutaraldehyde and embedded in Epon for autoradiographic exposure. I have also fixed with paraformaldehyde and embedded in Unicryl with polymerization by UV for immunogold labelling. I am mainly interested in bone and thus want to see if decalcification with EDTA is interfering with the immunogold label. Does anyone have any opinions as to which fixative/embedding resin would be best for this type of experiment?
TIA,
Pat Hales McGill University Dept. of Anatomy & Cell Biology hales-at-med.mcgill.ca
Return-Path: {Microscopy-request-at-sparc5.microscopy.com} Received: from rly-za05.mx.aol.com (rly-za05.mail.aol.com [172.31.36.101]) by air-za01.mail.aol.com (v51.9) with SMTP; Thu, 29 Oct 1998 09:41:11 -0500 Received: from Sparc5.Microscopy.Com (sparc5.microscopy.com [206.69.208.10]) by rly-za05.mx.aol.com (8.8.8/8.8.5/AOL-4.0.0) with SMTP id JAA11923; Thu, 29 Oct 1998 09:38:39 -0500 (EST) Received: (from daemon-at-localhost) by Sparc5.Microscopy.Com (8.6.11/8.6.11) id IAA21636 for dist-Microscopy; Thu, 29 Oct 1998 08:12:17 -0600 Received: from no_more_spam.com (Sparc5 [206.69.208.10]) by Sparc5.Microscopy.Com (8.6.11/8.6.11) with SMTP id IAA21633 for "MicroscopyFilteredEmail-at-msa.microscopy.com"; Thu, 29 Oct 1998 08:11:46 -0600 } From: "Mriglermas-at-aol.com"-at-sparc5.microscopy.com Received: from imo11.mx.aol.com (imo11.mx.aol.com [198.81.17.1]) by Sparc5.Microscopy.Com (8.6.11/8.6.11) with ESMTP id IAA21625 for {Microscopy-at-sparc5.microscopy.com} ; Thu, 29 Oct 1998 08:11:34 -0600 Received: from Mriglermas-at-aol.com by imo11.mx.aol.com (IMOv16.10) id NZJHa10441 for {Microscopy-at-sparc5.microscopy.com} ; Thu, 29 Oct 1998 09:21:34 -0500 (EST) Message-ID: {ea1bb27.363879ee-at-aol.com}
Our group currently has a JEOL FX 2000 TEM available for sale to anyone interested. The system includes a cold and hot stage, PC driven X-ray system, and STEM attachment. Please respond if interested.
M.W. Rigler, Ph.D. MAS, Inc. Suwanee GA 770-866-3218
Paula, For SEM studies of cells in suspension fixed with glutaraldehyde or frozen-freeze-substituted, we attach them to silane treated (1% silane from Aldrich in acetone for a few h in 50deg.C ) and carbon coated glass or mica chips. If they are conducitively stained, they do not require coating. The most important part of the procedure is to rinse off all the non-linked aldehyde groups from the cells and free silane from the surface of the chips. We have developed this procedure with Hans Ris for SEM studies of human blood normal and leukemic cells in which we were recovering ~95% of the cells. I will be happy to guide you through, provide more details, or literature (containing also descriptions of our attempts to use other methods) if within a sphere of your interests. Marek.
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It was recently stated on the list server that the Diatome Semi and the Histo are mostly the same, and that a Histo will do the same job as a Semi. This is not true at the TEM level. The semi knives are more highly polished and create a much smoother (higher resolution) surface on epoxides. The Diatome Semi is wonderful knife for thin sectioning because one can occasionaly take a one micron section to view (ultras can not be used like this). The Diatome Histo is a marvel for thick sections. We have 22 thousand dollars worth of Diatomes. We start students thin sectioning on a Histo with the understanding that these sections are not publication quality. I don't own stock in Diatome. Wish I owned the company! Bye, Hildy
Help! Are there any histology people out there who know the whereabouts of a German company called MED'LASS? We have a tissue processor made by them about 12-15 years ago which uses acetone to dehydrate and a vacuum chamber to infiltrate. It was called the Paraffinator. To my knowledge, there are only a handful of these instruments in the U.S. and I've been unsuccessful in trying to contact the manufacturer. ANY information would be helpful. Thanks, Bob Santoianni Emory University Hospital Atlanta, Georgia robert_santoianni-at-emory.org Tel. 404-712-4874
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On Fri, 30 Oct 1998, Alexander Mironov 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 } -----------------------------------------------------------------------. } =20 } =20 } Dear microscopists, } We are planning to buy some diamond knives and I would like to know your } impressions of knife performance (what is the difference between Diatome, } Drukker, Pelco, DDK etc.). We need standard ultramicrotomy knives for } cutting epon embedded biological material and cryo dry ultramicrotomy } knives. Personally, I like Diatome and dislike DDK. } Did anybody see the difference in cutting and durability between 45=B0 an= d } 35=B0 knives?=20 } Which boat for standard knife is preferable? What is the purpose of boats } with inclined cavity? } Any experience is wellcome. } =20 } ___________________________ } Dr. Alexander A. Mironov Jr. } Unit of Morphology } Dept. of Cell Biology and Oncology } Consorzio Mario Negri Sud } Via Nazionale, S.Maria Imbaro (Ch) } 66030 Italy } =20 } Tel. 0872-570-332 } Fax 0872-578-240 } E-mail: amironov-at-cmns.mnegri.it } =20 } =20 } =20 } =20 I have used DDK, Dupont, Diatome, others. For the last ten years we have invested only in Diatome. For the last seven years we have about 22 thousand dollars worth of assorted Diatome knives. We have Never had a bad one, never had one poorly resharpened (I long ago quit testing them when they came back to us), never had one that wore out quickly. Furthermore, Diatome USA has a laboratory set up. In case you have trouble with your embedding or materials, you can send them some of your tissues with your knife, and they will probe the situation for you and give you advice. I have not used this service, but I know from others that this service is fine. We have gotten excellent advice also on solving problems with our microtomes which was affecting the knives. I don't own stock in Diatome. Wish I did.
I'm interested in obtaining information about all commercially available cathodoluminescence detector systems. This includes both stand-alone and SEM add-on systems. I would like to generate a list of vendors that manufacture CL instrumentation.
Many thanks in advance. Vendors are welcome to contact me.
Best regards,
Angela
--------------------------------------------- Angela V. Klaus
Manager - Core Microscopy Facility American Museum of Natural History Central Park West at 79th Street New York, NY 10024-5192 USA
Hello. My recomandation for knifes would be to buy Drukker knives! It seems like most of you prefere the Diatome knife. At my lab we have during the last few years changed from Diatome to Drukker knifes, both for resin embedded material (Epon and Lowicryl)and for cryosections. All of us (we are seven people using this knives routinely) all think that the Drukker knives are better than Diatome, and for the two of us making cryosections most of the time it sure made sectioning easier.
I have a collaborator who wants to produce some EM samples using a = spraying procedure. The particles which are produced are quite large (up = to 50=B5m in diameter).=20
I know how to handle suspensions, but these particles are buoyant and so = will not settle. (Sorry to be so vague about details, but the work is = confidential in nature.)
I remember, from my undergrad EM courses, many years ago, that samples can = be sprayed using a =22nebulizer=22, but I can=27t seem to locate one to = try.
Can anyone help me locate a nebulizer, or suggest other options for = producing these samples?
As always, thanks for your help.=20
Paula.
Paula Allan-Wojtas Food Microstructure Specialist Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
--- --- --- -- -- -- --- --- --- Richard J. Dudley (rdudley+-at-pitt.edu) Research Specialist V Dept. of Cell Biology and Physiology University of Pittsburgh http://www.cbp.pitt.edu ---} search BIONET archives at http://www.bio.net {---
In another week or so I'll be leaving the art and science of scanning electron microscopy for a new career as a quality manager in our automotive lighting division. Before I go, I'd like to thank you, the worldwide community of microscopists in this forum, for your generous offerings of free advice, tips, and insights into our craft. You have allowed me to become a better microscopist and a better service provider for my customers. This is a debt I cannot repay other than to say that I will carry your spirit of openness and collaboration into my new job.
I'd also like to thank our friendly neighborhood system administrator for his uncanny ability to remain friendly in this neighborhood. I know I wouldn't have the patience to do what he does on our behalf.
Best wishes to all of you,
Harold J. Crossman OSRAM SYLVANIA INC. Lighting Research Center 71 Cherry Hill Dr. Beverly, MA 01915 (978) 750-1717 crossman-at-osi.sylvania.com http:///www.sylvania.com
Dear colleagues: We are recently asked to process mouse sperm for SEM. Do any of you have specific protocol for such specimen at hand? Or can we process the sample as ordinary cell suspension? Thanks in advance to any help. Yuhui Xu, MD EM Core, DFCI
I would like to thank all who shared their experience about the performance of diamond knives and told me the address of Agar. Thank you.
___________________________ Dr. Alexander A. Mironov Jr. Unit of Morphology Dept. of Cell Biology and Oncology Consorzio Mario Negri Sud Via Nazionale, S.Maria Imbaro (Ch) 66030 Italy
In a message dated 98-11-05 08:22:38 EST, allanwojtasp-at-em.agr.ca writes:
{ { I have a collaborator who wants to produce some EM samples using a spraying procedure. } }
Hi Paula,
I have used a nebulizer for just such purposes, but like you, I can't for the life of me remember where we got it. I recall vaguely that it was from a medical supply firm. The word "Vaponefrin" sticks in my mind, but this may be a trade name for an inhalable form of epinephrine rather than the name of the nebulizer.
But I have also used an artist's airbrush with good results. The airbrush is nice because you can regulate the air pressure and the size of the droplets which are produced. Plus, you can get one easily at your local arts and crafts supply store.
There are also stainless steel ultrasonic nebulizers which are hand-held devices (not the large ones for use in homes), but they tend to be quite expensive.
I would go with the airbrush if you can't find a nebulizer.
Good luck, hope this helps!
Cheers,
Bob **************************************** Robert (Bob) Chiovetti Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives / Systems Integrators / Analog & Digital Imaging *****************************************
We at Ladd Research, like many of our competitors, sell nebulizers. In our case it is catalog number 23625. If you are interested we could get you specs and a price.
Thabks,
JD Arnott
Paula Allan-Wojtas wrote: } } } Hi, all, } } I have a collaborator who wants to produce some EM samples using a spraying procedure. The particles which are produced are quite large (up to 50µm in diameter } } I know how to handle suspensions, but these particles are buoyant and so will not settle. (Sorry to be so vague about details, but the work is confidential in } } I remember, from my undergrad EM courses, many years ago, that samples can be sprayed using a "nebulizer", but I can't seem to locate one to try. } } Can anyone help me locate a nebulizer, or suggest other options for producing these samples? } } As always, thanks for your help. } } Paula. } } Paula Allan-Wojtas } Food Microstructure Specialist } Agriculture and Agri-Food Canada } Atlantic Food and Horticulture Research Centre } Kentville, Nova Scotia Canada B4N 1J5 } } Tel: (902) 679-5566 } FAX: (902) 679-2311 } } email: allanwojtasp-at-em.agr.ca }
--
LADD RESEARCH 13 Dorset Lane Williston, VT 05495
TEL 1-800-451-3406 (US) or 1-802-878-6711 (anywhere) FAX 1-802-878-8074 e-mail ladres-at-worldnet.att.net web site http://www.msa.microscopy.com/SM/LADD
We have used and inexpensive air-brush in the past. You can find them at= any hobby store and they are much more adjustable than a nebulizer. We = use N2 gas to operate the air-brush as it is cheap and basically inert. Greg
Paula Allan-Wojtas wrote:
} -----------------------------------------------------------------------= - } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Co= m } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.htm= l } -----------------------------------------------------------------------. } } Hi, all, } } I have a collaborator who wants to produce some EM samples using a spra= ying procedure. The particles which are produced are quite large (up to 5= 0=B5m in diameter). } } I know how to handle suspensions, but these particles are buoyant and s= o will not settle. (Sorry to be so vague about details, but the work is c= onfidential in nature.) } } I remember, from my undergrad EM courses, many years ago, that samples = can be sprayed using a "nebulizer", but I can't seem to locate one to try. } } Can anyone help me locate a nebulizer, or suggest other options for pro= ducing these samples? } } As always, thanks for your help. } } Paula. } } Paula Allan-Wojtas } Food Microstructure Specialist } Agriculture and Agri-Food Canada } Atlantic Food and Horticulture Research Centre } Kentville, Nova Scotia Canada B4N 1J5 } } Tel: (902) 679-5566 } FAX: (902) 679-2311 } } email: allanwojtasp-at-em.agr.ca
-- =3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D= =3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D= =3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D 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 =3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D= =3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D= =3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D=3D
Paula, 50 micron samples are a little large for EM. Why aren't you = using LM? Are these samples dry or already in liquid? Are they boyant due to = density or surface tension? Spraying samples or other techniques must be used = with caution as unwanted classification can be difficult to minimize = especially if your size distribution is large. Is your goal size distrubition? = Little can be suggested without knowing a lot more about the sample. Russ
-----Original Message----- } From: Paula Allan-Wojtas [mailto:allanwojtasp-at-em.agr.ca] Sent: Thursday, November 05, 1998 8:04 AM To: microscopy-at-sparc5.microscopy.com
Hi, all,
I have a collaborator who wants to produce some EM samples using a = spraying procedure. The particles which are produced are quite large (up to = 50=B5m in diameter).=20
I know how to handle suspensions, but these particles are buoyant and = so will not settle. (Sorry to be so vague about details, but the work is confidential in nature.)
I remember, from my undergrad EM courses, many years ago, that samples = can be sprayed using a "nebulizer", but I can't seem to locate one to try.
Can anyone help me locate a nebulizer, or suggest other options for producing these samples?
As always, thanks for your help.=20
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 process quite a bit of mouse sperm for SEM. I usually have the investigator wash it and put fixative (2% glutaraldehyde in cacodylate buffer) and then bring it to me. I let the cells fix for about two hours in the fridge. Then I poly-L-lysine (1%) coat 12mm round coverslips for about 10 minutes. I wash off the poly-L-lysine with distilled water, add a drop of sperm in fixative and let that sit for 10 minutes. Then I wash with 25% ethanol and dehydrate the coverslips to 100% ethanol. They can be critical point dried, coated and viewed in the sem. This works well for red blood cells too.
Lesley S. Bechtold Supervisor, Biological Imaging The Jackson Laboratory 600 Main St. Bar Harbor, ME 04609 207-288-6191
Well, we all know microscopy is expensive, but worth it. However, it is not always easy to convince the powers-that-be. Now we are being asked to carry some of the weight by doing commercial work. I seem to remember a thread about this some time ago and the problems of the University of Hawaii. Does anyone remember the problems? Does anyone at another university have any ideas about doing commercial work? Thanks for any help you might give me. Joyce Chicago State University
On Thu, 05 Nov 1998 08:04:26 Paula Allan-Wojtas wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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On Thu, 05 Nov 1998 08:04:26 Paula Allan-Wojtas 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
Paula. Ted Pella sells nebulizers. You could make one by putting a T in a stopper on a glass flask. Hook compressed gas to one side, and a pipette to the other; you need a small diameter tube to extend from the inside of the Pipette to the liquid in the flask. -be careful -a
On Thu, 05 Nov 1998 08:04:26 Paula Allan-Wojtas 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
Dear Paula, If you can produce the particles, just dry a drop or two of suspension onto a polished graphite-covered stub or a stub with a C-impregnated sticky tab and examine, or gold-coat and examine. A nebulizer is just any device that forces a liquid through an orifice to break it into small drops. The smaller the orifice or the greater the force, the smaller the drops. You wrote:
} Hi, all, } } I have a collaborator who wants to produce some EM samples using a spraying procedure. The particles which are produced are quite large (up to 50=B5m in diameter).=20 } } I know how to handle suspensions, but these particles are buoyant and so will not settle. (Sorry to be so vague about details, but the work is confidential in nature.) } } I remember, from my undergrad EM courses, many years ago, that samples can be sprayed using a "nebulizer", but I can't seem to locate one to try. } } Can anyone help me locate a nebulizer, or suggest other options for producing these samples? } } As always, thanks for your help.=20 } } Paula. } } Paula Allan-Wojtas
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 am setting a new TEM lab in my institute and need many things. One of them is carbon rod sharpener to make 3 mm diameter carbon rod with a cylindrical tip. I found that the commercial one was very expensive relatively. Could anybody suggest how I can get a generic sharpener at a low price? Any other suggestion is also appreciated.
Jondo Yun Department of Inorganic Materials Engineering Kyungnam University 449 Weolyeong-dong Masan, 631-701 Korea 82-551-249-2697 (office) 82-551-248-5033 (fax) 82-551-249-2692 (department office) 82-551-249-2719 (laboratory) 82-551-249-2564 (EM lab) email: jdyun-at-hanma.kyungnam.ac.kr
Be extremely cautious here. If any of your instruments were purchased using NSF funding and/or were purchased DUTY-FREE you will likely be violating various rules concerning the use of this equipment for commerical organizations. NSF has specific rules (NSF Notice 91) regarding instruments purchased with their funds (or at least they did last time I checked). Similiarly if you buy equipment duty free the forms (again the last time I filled one out) specifically require you to testify that it will not be used to be for commerical work for others.
There are alot of commerical "service" organizations that have paid full freight to buy their instruments and these RULES are made to protect their interests.
There is also a new law just passed in Congress S 314 which requires the governmental organizations (not defined as far as I know) to document any commerical activities and justify those which they perform. The intent appears to insure that the governmental resources do not compete with the commerical sector. It's a very narrow line and very easy to cross. I think at the moment that S314 is directed at Federal Employee's, but it could conceivably also be applied to anyone with Federal $$$.
I guess I'm waving a big red flag and telling you to tread carefully. In any event you had better point this out to your powers that be and have your legal eagles at CSU investigate all your potential liabilities. You could conceivably have to pay back the DUTY FREE fees or loose all future DUTY FREE purchasing ability.
The Santa Clara Valley Chapters of ASM and IEEE Reliability Society, with generous support from FEI Company, Micrion Corp, Schlumberger, and Nissei Sangyo America present:
NANOSCALE CHARACTERIZATION WITH THE FOCUSED ION BEAM Speaker: Prof. Robert Hull Department of Materials science and Engineering University of Virginia
ABSTRACT: Nanoscale characterization techniques using the focused ion beam (FIB) instrument will be reviewed. Dr. Hull will describe FIB-based techniques (either stand-alone, or in conjunction with transmission electron microscope imaging) for stress mapping in crystalline structures, dopant mapping in semiconductors, three dimensional image reconstruction, and ultra-high resolution secondary ion mass spectroscopy maps and volume reconstructions. He will also summarize additional techniques described in the literature, including FIB-induced optical emission spectroscopy and voltage contrast imaging. Finally specimen modification and damage artifacts created by the FIB beam will be discussed.
BIOGRAPHICAL SKETCH Robert Hull is an Associate Professor, and Doris and Heinz Wilsdorf Distinguished Research Chair, in the Department of Materials Science and Engineering at the University of Virginia. Prior to joining UVA he was a member of Technical Staff in the Physics Research Division of Bell Laboratories for seven years. He has authored and co-authored over one hundred and fifty papers in the fields of electronic materials, epitaxial growth, and applications of focused ion and electron beams. He has also given over fifty invited presentations at national and international conferences in these fields. He is on the editorial board of several major journals, and has edited several proceedings and reference volumes. In 1997 he was the President of the Materials Research Society, the leading international society in the field of materials science and engineering.
TIME AND LOCATION: November 11 at David's Restaurant at the Santa Clara Tennis and Golf Club, at 5151 Stars & Stripes Drive in Santa Clara, CA (95054). This is just east of the Santa Clara Convention Center. Dinner choice of: London Broil or Seafood Brochette served with lemon butter sauce or a Vegetarian (Pasta Primavera) plate. Social at 6:00 p.m., 6:45 Dinner and 8:00 p.m. Talk
Cost: ASM/IEEE Members $16, Students $8 and Guests $18 (with an additional two dollars if no RSVP the Monday before the event (i.e, Nov 9th) Reservations: Brock Hinzmann 650-859-4350 or email IEEE Santa Clara Valley Chapter Reliability Society contact David Su (davidsu-at-aol.com). Please include choice of meal!
At 09:51 05.11.98 EST, you wrote: } What is it that you base your decision on. Do you get a kickback from } Drukker? }
No kickback from Drukker! But I'm one of those who actually have had the oprtunity to compare the kniver, and like the Drukker ones. If that is not good enough I can't see the value of asking about peoples opinions! By the way; who is actually behind the SGKCCK adress??? RO
Randi Olsen Department of Electron Microscopy Faculty of Medicine University of Tromso MH-Breivika N-9037 TROMSO NORWAY
Dear all, I am passing this message on for a colleague who is wondering about purchasing a better scanner, specifically for TEM plate negatives. He already has a UMAX Astra 1200S with transparency adaptor but this does not cope with high optical densities, or with enlarging small areas greatly. We use two different sizes of plate film: one just smaller than 3 1/4" by 4" and the other 2 1/2" by 3 1/2". What would you recommend, a high end flatbed with transparency adaptor or a specialist negative scanner? Which models have worked well for you. He would prefer it to be Mac compatible.
Thanks
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ 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. ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
We have dealt with this issue extensively. You can always try to do this through your office of sponsored programs or wahatever they call it there. The outside person can give you a grant or gift to do the work. We then put the money in a University Foundation Account. You may also be able to enter into a contract for research or services . What if any overhead the university might take needs to be negotiated. We generally no longer operate in that manner since we have been reorganized as an auxiliary business. That is we recharge our internal customers and are able to serve outside cutomers as resources permit. This is a business incidental to an educational activity and is looked on much as the university bookstore is. Laws and regulations vary from state to state. Fell free to contact me for more info. I also got into Federal Cost Accounting morass quite deeply and can discuss that with you as well.
Greg At 03:19 PM 11/05/1998 -0600, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Thanks for all of you who responded with locations for nebulizers, and other options for spraying the samples. Some we have tried already, and some we will try within the next month.
I can give you a little more information, and ask for help with a slightly different problem this time. We are also looking for ways of producing the sample, not just depositing it on EM supports (grid or stub). The collaborator tells me that they are now able to make the samples quite abit smaller in diameter using thier traditional methods.
The reason why we are using EM at all is that we are studying the microstructure of the sample and changes in it. The collaborator uses LM as a quick check after each modification is made.
I remember seeing a method where a grid or some support is placed in a centrifuge tube with the sample, and the sample is "spun" onto the support. Does anyone have a reference for this one?
Thanks very, very much!
Paula.
Paula Allan-Wojtas Food Microstructure Specialist Agriculture and Agri-Food Canada Atlantic Food and Horticulture Research Centre Kentville, Nova Scotia Canada B4N 1J5
1) histo knives - you are only partly correct, Hildegard. In terms of materials science 'hard' materials sectioning, histo knives can indeed furnish publication quality micrographs in the sense that the sectioning process itself inevitably induces so much deformation damage in our crystalline materials that the 'rippled' surface produced by the histo is of little consequence relative to the desired information from the section. (I call it 'continuous knife marks' because that is what it looks like). No less a personage than Prof. Helmut Sitte in Germany agrees with our thinking that this may be due to a serrated nature for the knife edge that might just serve the purpose of a self-sharpening effect of sorts. We have tested histo knives from both Diatome and DDK on metals (some quite hard) and have observed the following (summarized also in EMSA/92, p. 294): - they produce quite good ultrathin sections (if one ignores the above marks), indeed the thinnest sections we have ever produced - {10nm in Al - were with a Diatome histo, - they produce incredibly flat sections, seemingly because the unique nature of the edge somehow counteracts the asymettric buildup of deformation that produces the dreaded curling of metallic sections (like in lathe turnings) that drives our microtomist nuts at times, - for some ultrahard metals (FE-Nd-B amorphous intermetallic particles), it was the only knife (compared to 35, 45 and 55 degree colleagues) that produced useable sections, - before you all rush out and buy histos for materials science sectioning, they also; vary quite a bit more in performance than regular knives (what do you expect for half the price?), may start to 'shred' the section as the thickness drops below 100nm, and are hopeless for cutting anything with weakly adherent interfaces (coatings and the like).
2) As per the related thread about Diatome vs Drukker, etc; we have used knives from Diatome, Drukker, Microstar and DDK over the last 15 years (all for materials science sectioning), yet I am always loathe to give any sort of public ranking or recommendations. A lot depends on the materials you are sectioning, your relationship with the knife producer, how you clean and care for your knives, what is the stage the edge at any given time, etc. Our person prefers the Diatomes (we have several), but has obtained good results from all of the others, so-----, each to your own. The important point is that, over the years, I have found most diamond knife producers to be amazingly cooperative and helpful with regard to performance questions, tips and hints, resharpening and the like. If only some of the other makers of EM accessories were so helpful!
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: HILDEGARD CROWLEY[SMTP:hcrowley-at-du.edu] } Sent: November 04, 1998 2:02 PM } To: postmessage } Subject: Re: Diatome Semi, Ultra, Histo } } ------------------------------------------------------------------------ } 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, } } It was recently stated on the list server that the Diatome Semi and the } Histo are mostly the same, and that a Histo will do the same job as a } Semi. This is not true at the TEM level. The semi knives are more highly } polished and create a much smoother (higher resolution) surface on } epoxides. The Diatome Semi is wonderful knife for thin sectioning because } one can occasionaly take a one micron section to view (ultras can not be } used like this). The Diatome Histo is a marvel for thick sections. We } have 22 thousand dollars worth of Diatomes. We start students thin } sectioning on a Histo with the understanding that these sections are not } publication quality. } I don't own stock in Diatome. Wish I owned the company! } Bye, } Hildy } }
Could you please send this to the EM Server? Thanks.
We are looking at EM tissue processors and hope to make a purchase soon. We process primarily biological samples. Does anyone have experience with the EMP5160 manufactured by RMC? It is similar to the LKB tissue processor that we used to have but seems to have some nice modifications. If anyone has experience with this model (pros or cons), or any other suggestions, please contact me at gatlin_cindy_l-at-lilly.com or reply via server if you think others might be interested.
Thanks very much,
Cindy Gatlin Technical Associate/EM lab Eli Lilly & Co.
Ian asks ... } } } Dear all, } I am passing this message on for a colleague who is wondering about } purchasing a better scanner, specifically for TEM plate negatives. He } already has a UMAX Astra 1200S with transparency adaptor but } this does not } cope with high optical densities, or with enlarging small } areas greatly. } We use two different sizes of plate film: one just smaller } than 3 1/4" by 4" and the other 2 1/2" by 3 1/2". } What would you recommend, a high end flatbed with } transparency adaptor or a specialist negative scanner? ...
Since EM image acquisitions typically are the result of "scans" and therefore depict scan lines, I might suspect a scanner for specific film types might cause moire patterns. That is, the scanner would insist you mount the film almost orthogonally, whereas you might try to remedy moire patterns by tilting the film on a higher res flatbed ... say, 30deg, and then making it orthogonal again with your favorite image editor. Dedicated film scanners would offer the resolution you want ... I'd simply suggest you try them before you buy them.
... 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/
} Well, we all know microscopy is expensive, but worth it. However, it is } not always easy to convince the powers-that-be. Now we are being asked } to carry some of the weight by doing commercial work. I seem to } remember a thread about this some time ago and the problems of the } University of Hawaii. Does anyone remember the problems? Does anyone } at another university have any ideas about doing commercial work? } Thanks for any help you might give me. } Joyce } Chicago State University
Joyce -
The problem in Hawaii was different; it had to do with the way lab usage was being charged to federal grants. I dealt with commercial usage pre-retirement at U.C. Berkeley. There were very specific statewide university rules that covered all such use on the campus. I was allowed to provide service to outsiders only if there was no private lab in the local area offering the same service, and I was told to charge double the campus recharge rate. I supported the policy, because competition with private enterprise using grant-funded equipment really is unfair.
Caroline
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
When we decided to buy a scanner for negatives we tested different scanners, from the top-end scanner used for cartography (extremely expensive and with pixel size of about 7.5 microns) down to flatbed scanners with transparency adaptor. From our tests we obtained that for TEM plates negative scanners were better and that for high resolution TEM work at least (optical) 1600 dpi were required in order to be able to enlarge the image sufficiently. We decided to buy a Polaroid SprintScan 45, which accepts large negatives. We use 2000x2000 dpi for HREM work. Optical density is about 3.4, if I'm not wrong. We are satisfied with this instrument, except for the negative holder: for 3 1/4"x4" negatives the holder is too large and some bending of the negative occurs, as it can only be held on the two short sides. However we didn't notice large image distortion. I think that I read in this list that Polaroid was preparing special holders for the negatives, but I don't know further details. Is there somebody that could answer this question? For smaller negatives an adapter exist with two magnets, but I don't know whether your negatives can be fitted in it (included with the scanner). As conclusion, during the last year I didn't use the dark room at all and I think that I will extremely seldomly use it again (only if our dye-sub printer breaks). I hope this helps.
Albert Albert Romano-Rodriguez Dept. of Electronics Faculty of Physics University of Barcelona c/ Marti i Franques, 1 E-08028 BARCELONA Spain tel: +34-93-402 11 47 FAX: +34-93-402 11 48 e-mail: romano-at-el.ub.es
We are in the early stages of evaluating a digital imaging system here (for light microscopy). I'm trying to understand the different ways that CCD cameras can be or are used to acquire color images. There seem to be several ways this is done:
* Single chip Monochrome CCD array with some type of filter in front of the camera to allow it to acquire sequential red, green & blue images and then software to "reassemble" the images into a color image. * 3 Chip camera, with each chip assigned (filtered for) red, green & blue and then software to "reassemble" the images into a color image. * Single chip CCD with some type of color mosaic "mask" on the chip to acquire the red, green & blue parts of the image and then software to "reassemble" the images into a color image. * CCD array where the image is "scanned" either by moving optical elements or moving the CCD array to acquire a high pixel count with a fairly small sensor. These would be the slowest for acquisition, but I gather they give a lot of "bang for the buck (euro)". * Others?
What are the Pros & Cons of these different types of cameras? Is there a WWW or published resource that could help me sort this out?
Our interest here is primarily in acquiring static images (not real-time video) from low light level fluorescence, DIC and/or bright field. We would like the camera to be sensitive enough for quantitation if needed. I'd also like to be able to advise others here who may have different requirements. I'm not specifically "fishing" for sales pitches (I already have plenty of glossy literature, I'm just trying to make sense out of it).
I would be happy to take replys "off-list" and post a summary.
Yours, Doug .................................................................... : Douglas W. Cromey, M.S. Dept. of Cell Biology & Anatomy : : Research Specialist, Principal University of Arizona : : (office: AHSC 4212A) P.O. Box 245044 : : (voice: 520-626-2824) Tucson, AZ 85724-5044 USA : : (FAX: 520-626-2097) (email: doug-cromey-at-ns.arizona.edu): :...................................................................: http://www.pharmacy.arizona.edu/exp_path.html Home of: "Microscopy and Imaging Resources on the WWW"
} I am passing this message on for a colleague who is wondering about } purchasing a better scanner, specifically for TEM plate negatives. He } already has a UMAX Astra 1200S with transparency adaptor but this does not } cope with high optical densities, or with enlarging small areas greatly. } We use two different sizes of plate film: one just smaller than 3 1/4" by } 4" and the other 2 1/2" by 3 1/2". } What would you recommend, a high end flatbed with transparency adaptor or a } specialist negative scanner? Which models have worked well for you. He } would prefer it to be Mac compatible. } Dear Ian, I think the best scanners are either the flat-bed (like the Perkin Elmer) or rotating Drum (like the Optronics). These are capable of 5-by-5 micrometer pixel size, in the case of the PE, and the files produced can by presented in any size needed depending on the image processing program used. The SPIDER system used here will handle both PE and Op data. The bad news is that these are running on a mainframe, and, AFAIK, not Mac compatable. There may be software out there which runs on the Mac and can read PE or Op files. Good luck. Yours, Bill Tivol
=C0=B1 =C1=B8=B5=B5 Jondo Yun wrote: } =20 } -----------------------------------------------------------------------= - } The Microscopy ListServer -- Sponsor: The Microscopy Society of Dear Jo= ndo, =20 } I am setting a new TEM lab in my institute and need many things. } One of them is carbon rod sharpener to make 3 mm diameter carbon rod wi= th a } cylindrical tip. I found that the commercial one was very expensive } relatively. } Could anybody suggest how I can get a generic sharpener at a low price? } Any other suggestion is also appreciated. } =20 Ted Pella sells a hand-held sharpener for under $100. You should check his web site for info. I am not connected with Pella except as a customer. Yours, Bill Tivol
Michael Shaffer's comment seems a bit off the mark. TEM negatives are generally not scanned images. Although some types of images (e.g., crystal lattice images) can give moire's when scanned, this effect can be avoided by appropriate choice of scanner sampling resolution. I concur with the 'try before you buy' philosophy, if the opportunity can be made.
The optical density range of high-end 12-bit grayscale scanners (36 bit color) goes as high as 3.6. Unfortunately, scanner software seldom offers exposure control. Actually, 3.6 or even (more common) 3.2 is an adequate density range for most TEM photos including diffraction patterns. Your microscopist colleague might consider controlling exposures in the microscope to avoid excessively dense negatives. There are also special film developing solutions that might be used for special situations where contrast reduction is necessary.
My personal observation of current 36-bit, } 1000 spi scanners is that they are incredibly slow and inefficient for scanning TEM negatives. Anyone contemplating a market survey should include in their evaluation the time takes to scan into Photoshop including time the scanner takes calibrating itself before each scan.
Don't know if you saw my earlier posting to the listserver concerning scanning TEM negatives. The essence was that the 'negative film scanning mode' gives poor grayscale results for high-contrast TEM negatives because of a built in contrast correction feature. The scanned images become posterized. The work-around is to always scan in the positive transparency mode and invert the image contrast with other controls.
Larry Thomas Mechanical and Materials Engineering Washington State University Pullman, WA USA
---------- From: shAf Sent: Friday, November 6, 1998 5:01 PM To: Ian MacLaren; Microscopy-at-Sparc5.Microscopy.Com Cc: g.r.millward-at-BHAM.AC.UK Subject: RE: Scanning negatives
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Ian asks ... } } } Dear all, } I am passing this message on for a colleague who is wondering about } purchasing a better scanner, specifically for TEM plate negatives. He } already has a UMAX Astra 1200S with transparency adaptor but } this does not } cope with high optical densities, or with enlarging small } areas greatly. } We use two different sizes of plate film: one just smaller } than 3 1/4" by 4" and the other 2 1/2" by 3 1/2". } What would you recommend, a high end flatbed with } transparency adaptor or a specialist negative scanner? ...
Since EM image acquisitions typically are the result of "scans" and therefore depict scan lines, I might suspect a scanner for specific film types might cause moire patterns. That is, the scanner would insist you mount the film almost orthogonally, whereas you might try to remedy moire patterns by tilting the film on a higher res flatbed ... say, 30deg, and then making it orthogonal again with your favorite image editor. Dedicated film scanners would offer the resolution you want ... I'd simply suggest you try them before you buy them.
... 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/
Nestor's caution is fully justified. His description is right on the mark, and the rules have just been re-issued as Important Notice number 122 of 1998 (see http://www.nsf.gov/pubs/1998/iin122/iin122.txt).
Caroline was generous. We charge commercial customers three times our academic rate - but the academic rate is the same for MIT and any other academic user.
The microscope is currently in use. It has some good days and some bad days and it will be available around Thanksgiving. It's free!! (If your willing to haul it away.) The SEM resides in Easton PA.
If you think you might be interested, give me a call: Nan Laudenslager Specialty Minerals, Inc. (610) 250-3094
} Some diamond knife comments: } } 1) histo knives - you are only partly correct, Hildegard. In terms of } materials science 'hard' materials sectioning, histo knives can indeed } furnish publication quality micrographs in the sense that the sectioning } process itself inevitably induces so much deformation damage in our } crystalline materials that the 'rippled' surface produced by the histo is of } little consequence relative to the desired information from the section. (I } call it 'continuous knife marks' because that is what it looks like). No } less a personage than Prof. Helmut Sitte in Germany agrees with our thinking } that this may be due to a serrated nature for the knife edge that might just } serve the purpose of a self-sharpening effect of sorts. We have tested } histo knives from both Diatome and DDK on metals (some quite hard) and have } observed the following (summarized also in EMSA/92, p. 294): } - they produce quite good ultrathin sections (if one ignores the above } marks), indeed the thinnest sections we have ever produced - {10nm in Al - } were with a Diatome histo, } - they produce incredibly flat sections, seemingly because the unique nature } of the edge somehow counteracts the asymettric buildup of deformation that } produces the dreaded curling of metallic sections (like in lathe turnings) } that drives our microtomist nuts at times, } - for some ultrahard metals (FE-Nd-B amorphous intermetallic particles), it } was the only knife (compared to 35, 45 and 55 degree colleagues) that } produced useable sections, } - before you all rush out and buy histos for materials science sectioning, } they also; vary quite a bit more in performance than regular knives (what do } you expect for half the price?), may start to 'shred' the section as the } thickness drops below 100nm, and are hopeless for cutting anything with } weakly adherent interfaces (coatings and the like). } } 2) As per the related thread about Diatome vs Drukker, etc; we have used } knives from Diatome, Drukker, Microstar and DDK over the last 15 years (all } for materials science sectioning), yet I am always loathe to give any sort } of public ranking or recommendations. A lot depends on the materials you } are sectioning, your relationship with the knife producer, how you clean and } care for your knives, what is the stage the edge at any given time, etc. } Our person prefers the Diatomes (we have several), but has obtained good } results from all of the others, so-----, each to your own. The important } point is that, over the years, I have found most diamond knife producers to } be amazingly cooperative and helpful with regard to performance questions, } tips and hints, resharpening and the like. If only some of the other makers } of EM accessories were so helpful! } } } 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: HILDEGARD CROWLEY[SMTP:hcrowley-at-du.edu] } } Sent: November 04, 1998 2:02 PM } } To: postmessage } } Subject: Re: Diatome Semi, Ultra, Histo } } } } ------------------------------------------------------------------------ } } 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, } } } } It was recently stated on the list server that the Diatome Semi and the } } Histo are mostly the same, and that a Histo will do the same job as a } } Semi. This is not true at the TEM level. The semi knives are more highly } } polished and create a much smoother (higher resolution) surface on } } epoxides. The Diatome Semi is wonderful knife for thin sectioning because } } one can occasionaly take a one micron section to view (ultras can not be } } used like this). The Diatome Histo is a marvel for thick sections. We } } have 22 thousand dollars worth of Diatomes. We start students thin } } sectioning on a Histo with the understanding that these sections are not } } publication quality. } } I don't own stock in Diatome. Wish I owned the company! } } Bye, } } Hildy } } } } } I have never been involved with materials science. My remarks only apply to biological materials.
DVC Company has an answer to your need for a decent high resolution camera that others reading this might need. DVC has introduced this at the Vision Show in San Jose last month and at the Photonics East in Boston last week and now.....at the Neuroscience show in LA at our DVC booth in the 8XX area starting this Sunday. DVC does the whole system and the color info is done in the host/ 400Mhz computer which we supply along with the Epix frame grabber that is only $995.
See our web http://members.aol.com/dvcco
We have a 1300 x 1030 pixel digital camera that uses a single sensor with a color Bayer pattern. The singel sensor offers in effect } 700 TV Lines square from our model DVC1300C and in monochrome at } 1000 TV Lines Square. Using a single sensor offering 12 frames / sec is much better if real time 30fps is not needed because we offer the first camera to do this with 58-60dB S/N !!! You have no color shift as with 3 chip CCD cameras with a price of $4995. You also have a higher vertical resolution as stated above / square pixels, and no problem with ( color shift ) which comes from misalighnment of the 3 chips on a prism situation and the cost......10K plus....and bulky also. Our unit weigh's less than one pound. You can go with a bulky expensive $1500-2500 attachable LCD filter on the monochrome version of the camera but........you loose 30% sensitivity.....and only gain perhaps 200 TV Lines over our 700TV Line unit.... You should check out our web on this and visit us at Neuroscience. For those who feel offended from learning something new, especially from the camera manufacturer..... well excuse me ! With all due respect for the others that do not. Regards,
Rich
Richard Klotsche DVC Company/ San Diego 619-444-8300 619-444-8321-fax
In a message dated 98-11-06 14:17:40 EST, you write:
{ { Subj: Color CCD Camera ?s Date: 98-11-06 14:17:40 EST From: doug-cromey-at-ns.arizona.edu (Doug Cromey) To:
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Microscopy Listers,
We are in the early stages of evaluating a digital imaging system here (for light microscopy). I'm trying to understand the different ways that CCD cameras can be or are used to acquire color images. There seem to be several ways this is done:
* Single chip Monochrome CCD array with some type of filter in front of the camera to allow it to acquire sequential red, green & blue images and then software to "reassemble" the images into a color image. * 3 Chip camera, with each chip assigned (filtered for) red, green & blue and then software to "reassemble" the images into a color image. * Single chip CCD with some type of color mosaic "mask" on the chip to acquire the red, green & blue parts of the image and then software to "reassemble" the images into a color image. * CCD array where the image is "scanned" either by moving optical elements or moving the CCD array to acquire a high pixel count with a fairly small sensor. These would be the slowest for acquisition, but I gather they give a lot of "bang for the buck (euro)". * Others?
What are the Pros & Cons of these different types of cameras? Is there a WWW or published resource that could help me sort this out?
Our interest here is primarily in acquiring static images (not real-time video) from low light level fluorescence, DIC and/or bright field. We would like the camera to be sensitive enough for quantitation if needed. I'd also like to be able to advise others here who may have different requirements. I'm not specifically "fishing" for sales pitches (I already have plenty of glossy literature, I'm just trying to make sense out of it).
I would be happy to take replys "off-list" and post a summary.
Yours, Doug .................................................................... : Douglas W. Cromey, M.S. Dept. of Cell Biology & Anatomy : : Research Specialist, Principal University of Arizona : : (office: AHSC 4212A) P.O. Box 245044 : : (voice: 520-626-2824) Tucson, AZ 85724-5044 USA : : (FAX: 520-626-2097) (email: doug-cromey-at-ns.arizona.edu): :...................................................................: http://www.pharmacy.arizona.edu/exp_path.html Home of: "Microscopy and Imaging Resources on the WWW"
We have been using an RMC tissue processor 5160 for several years. Our processor may be a bit different than the one currently sold, as it not only is designed to keep the reagent vial in which the specimens reside at a specific temperature, it will also pre-cool the solution vial in which the samples are about to be placed as well. I believe that the newer model eliminated temperature control on the "next" vial, which in my mind means that all the solutions should be used at ambient temperature to avoid significant temperature fluctuation within the samples.
There are several "quirks" with the use of the machine which you will learn over time (or contact me to avoid reinventing the wheel). For example, we do not use the reagent vial caps unless necessary, since sometimes the caps are not secured to the vials during refitting (an alignment problem). The caps then fall into the mechanics and cause a jam. We also put small holes in the caps above volatile solutions (propylene oxide, 100% ethanol, acetone) since evaporation will cause the lids to lift up and become askew, causing an alignment problem which again causes the caps to fall into the machinery.
We have had some difficulties obtaining expendable, including specimen holders and particularly reagent vials. The design of the vials has been improved over time (they have thicker walls and don't warp as easily...warping caused the specimens to get stuck as they were transferred from one vial to the next, eventually causing the samples to be lost or confused with others).
Overall, despite the above comments, I really like the processor. Although it takes about 30 minutes to set up, it eliminates countless hours spent in the fume hood transferring solutions. Exposure to organic solvents, carcinogens and fixatives is much less likely. The machine allows overnight processing of tissues. We reuse many of our reagent vials (not the ones containing epoxies or OsO4) so it is not too expensive to use. It does use more volume of reagents than you may be accustomed to. We routinely use 10 to 18 ml in each of the 22 reagent vials, depending on how many samples we are processing (up to 12). Although I have done so, I am nervous to use the processor for tissues which are irreplaceable. If I do use it for these tissues, I prefer to do so when either myself or my technician is available to visit the machine occasionally to observe its performance.
I'd be happy to talk more with you over the phone.
Best wishes,
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 optical density range of high-end 12-bit grayscale scanners (36 bit color) } goes as high as 3.6. Unfortunately, scanner software seldom offers exposure } control.
While I concur with you about most of your message, I am surprised by this statement. We have several scanners on our floor and all of them have software which permits control of density and contrast, both manually and via presets. One can shift the grayscale 'window'quite easily and extract information even from quite dense negatives.
Ian, I compared some scanners for TEM negatives a few months ago. It all depends on the level of quality needed and how much enlarging you mean . 1200 dpi was sufficient for low mag. stereology and morphometry, and contrasty high mag. such as colloidal gold and dense histochemical deposits. At 2400 dpi the film's silver grains could be distinguished at high enlargements, suggesting nearly lossless scan. The 2400 dpi scans were suitable for low contrast, high mag. work and enlargments. The scanner everyone really loved was the Imascan, by Imacon (www.imacon.dk). It is essentially a small drum scanner with a flexible magnetic holder for positives or negatives that is very easy to use. At 5760 dpi, 14 bit and a Dmax of of 4.1, the images it produced could be enlarged to give distinct view of silver grains. These files did amount to about 30 Mb for a monochrome 3 1/4x4 negative. I put some extremely dense negatives through it with excellent results. the price on the Imascan is probably steeper than you want, US$16,000. The main EM users in our group are contemplating a joint purchase. The Imascan is driven by its own software for Mac or Windows and connects to either platform via SCSI port. I tested it with a 233 MHz G3 and found the scan times tolerable unless one was doing RGB at 2400 dpi or higher. then the 3 passes added up. 2 other scanners we are considering as an alternative to the Imascan are the LinoColor Ultra Sapphir and the Agfa Duoscan. They seem to be better able to get closer to their stated Dmax than some of the other brands.
Glen
Glen MacDonald Research Scientist Hearing Research Laboratories of the Virginia Merrill Bloedel Hearing Research Center Box 35-7923 University of Washington Seattle, WA 98195-7923 (206) 616-4156 glenmac-at-u.washington.edu
On Fri, 6 Nov 1998, Ian MacLaren wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Dear all, } I am passing this message on for a colleague who is wondering about } purchasing a better scanner, specifically for TEM plate negatives. He } already has a UMAX Astra 1200S with transparency adaptor but this does not } cope with high optical densities, or with enlarging small areas greatly. } We use two different sizes of plate film: one just smaller than 3 1/4" by } 4" and the other 2 1/2" by 3 1/2". } What would you recommend, a high end flatbed with transparency adaptor or a } specialist negative scanner? Which models have worked well for you. He } would prefer it to be Mac compatible. } } Thanks } } ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ } 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. } ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ } } } }
Hi Doug, I'm in almost the same situation that you describe. Can you post a summary once you have assembled replies to these questions? Here are a couple of URL's that I used when I was searching the web for info. Caution though, I got really bogged down. As you say, lots of glossy brochures but it's difficult to read between the lines (note the resolution pun) when you are limited on experience. Good luck. John
I've had good results using a Polaroid SprintScan45. They gave me a steel plate adapter with magnetic strips to insert in the 4x5 negative carrier and this worked fairly well. I went a step further and had a replacement plate made for TEM negatives that replaces the 4x5 spring loaded plate that comes with this holder. It works fairly well. The scanner is reasonably priced. Check out Polaroid's web site for specifications. -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: Ian MacLaren To: Microscopy-at-Sparc5.Microscopy.Com Cc: g.r.millward-at-BHAM.AC.UK -----------------------------------------------------------------------.
Dear all, I am passing this message on for a colleague who is wondering about purchasing a better scanner, specifically for TEM plate negatives. He already has a UMAX Astra 1200S with transparency adaptor but this does not cope with high optical densities, or with enlarging small areas greatly. We use two different sizes of plate film: one just smaller than 3 1/4" by 4" and the other 2 1/2" by 3 1/2". What would you recommend, a high end flatbed with transparency adaptor or a specialist negative scanner? Which models have worked well for you. He would prefer it to be Mac compatible.
Thanks
++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++ 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. ++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++++
Our group currently has a JEOL FX 2000 TEM available for sale to anyone interested. The system includes a cold and hot stage, PC driven X-ray system, and STEM attachment. Please respond if interested.
M.W. Rigler, Ph.D. MAS, Inc. Suwanee GA 770-866-3218
} Nestor's caution is fully justified. His description is right on the mark, } and the rules have just been re-issued as Important Notice number 122 of } 1998 (see http://www.nsf.gov/pubs/1998/iin122/iin122.txt). } } Caroline was generous. We charge commercial customers three times our } academic rate - but the academic rate is the same for MIT and any other } academic user. } } Tony. } I've gotten similar comments offlist; apparantly I didn't make my point clearly. Two times academic rates was a university-wide policy, and I had no authority to change it (tho I wanted to). Anyone setting up a new system needs to check on the existence of similar rules.
Caroline
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 am looking for a source to buy a Durst Laborator enlarger (with a point source and variable condensors) for printing our TEM negatives. I have contacted several sources on the web, including Photoshopper and Durst ACS, Inc., and have been unsuccessful in eliciting ANY responses from ANY vendor for purchasing a new Durst enlarger. I understand from my local photog supplies vendor that Durst is a quirky company with only a few authorized reps in the U.S.
Can anyone help me find a Durst vendor?
Offline replies would be fine. If others express interest, I will publish what I learn. Thanks to all.
Ann Hein Lehman TEM Lab Mgr, Trinity College LSC 314 300 Summit St. Hartford, CT 06106 860-297-4289 Ann.Lehman-at-exchange.cc.trincoll.edu
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Good luck. I had a helluva time getting any info out of their Midwest representative (some camera shop in Milwaukee) several years ago. I don't see how they stay in business. However, if you do find someone who will actually talk to you, please send me their name. I still have some questions about the finer details of operation. On the other hand, maybe with you could pick up a good used enlarger from a lab going digital.
The MSA office received a request from a high school recently for a brochure on "careers in microscopy". MSA doesn't have one, but I suspect that some of you who read this list may. If you have one, will you please send me a sample so that MSA and MICRO can refer future inquiries?
Caroline
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
Soon after I asked the question about the ion-milling with a cooled stage, I received a great deal of information concering this problem. Now, we solve it. Many thanks for your kind suggestion and help.
Yours Sincerely,
J.S. Wu --------------------------------- Jinsong WU LSG2M, Ecole des Mines de Nancy Parc de Saurupt F-54042 NANCY cedex France
Caroline Schooley wrote: ============================================= The problem in Hawaii was different; it had to do with the way lab usage was being charged to federal grants. I dealt with commercial usage pre- retirement at U.C. Berkeley. There were very specific statewide university rules that covered all such use on the campus. I was allowed to provide service to outsiders only if there was no private lab in the local area offering the same service, and I was told to charge double the campus recharge rate. I supported the policy, because competition with private enterprise using grant-funded equipment really is unfair. =================================================== Thanks Caroline, I am sure that I can speak for the many others performing electron and light microscopy as a for-profit tax-paying laboratory service wishing that the idea of this kind of "unfairness" was more universally recognized (and appreciated).
However, more often than not, such policies are formulated in ways that, like no-fault automobile insurance in many states, sound like they are accomplishing something of significance but in the end the life goes on for the lawyers as if nothing had changed.
} From my own almost thirty years of experience with this issue, from what I have seen, one would have to multiply the internal charge rate, in many instances, by factors of 5X to 10X before the "selling price" to the client would be truly "competitive" with those of commercial firms offering comparable services.
Second, the idea of there being "no private lab in the local area offering the same service," does not work either. For one thing, how does one define "local"? And how does one determine if it is the "same service"? We recently lost what for us would have been a nice SEM job, it was a large number of repetitive samples of glass spheres, highest magnification being about 1000X, to a university that justified their being able to do the work because they had an FE-SEM and we had a more plebeian tungsten filament SEM. And of course, the client had to drive about five miles further to get to us than to the university. So those those who manage the laboratory, and who have the most to benefit either via the normal structure of rewards present in such settings or via outright consulting fees, are also the ones making the decisions as to what is "local" and what is the "same". Sort of seems like it is the fox guarding the chicken coop (at least in some instances).
NSF Important Notice 122, like with the no fault insurance laws have the appearance like it is accomplishing something but those of us on the firing line on a daily basis, having to compete in the marketplace with university competitors, know that in most instances such policies are just window dressing and don't, at the end of the day, accomplish the stated goals.
Chuck
=================================================== Charles A. Garber, Ph. D. Ph: 1-(610)-436-5400 President 1-(800)-2424-SPI Structure Probe, Inc. 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 have a student attempting to immunogold label G-protein in arabidopsis guard cells. The antibody is made to aribidopsis protein expressed in E. coli. We can easily label E. coli cells expressing the protein and the antibodies recognize the protein purified from plants and run on westerns. No light level work has been done. However we are having a number of problems attempting to label the protein in Arabidopsis: the ultrastructure is terrible, and we have not even a hint of label in the plants. Any advice would be appreciated.
I would also appreciate advice on how to handle Arabidopsis guard cells for EM. We are trying leaf peels and diced leaves but both are difficult to consistantly section and get good guard cells. The student attempting this project, and doing all the embedding and sectioning, is an undergraduate; so any surefire advice you can offer for sectioning guard cells would help.
Thanks in advance,
Michelle
#################################################### Michelle Peiffer Electron Microscope Facility for the Life Sciences The Biotechnology Institute for Research and Education 1 South Frear Lab University Park, PA 16802 814-865-0212 email:mlk101-at-psu.edu ####################################################
I think we need some additional information on the nature of the Arabidopsis protein and how you are doing your specimen prep before we could make any recommendations.
Since you can label it in E. coli and it shows up OK on blots, it sounds like the antibody is doing what it's supposed to do. Since I'm not a botanist, I will have to ask a basic question: what is the nature of the protein in native guard cells? Is it water soluble? Hydrophobic? A membrane protein?
You mention that the ultrastructure is terrible, and this may be the root of the problem (no botanical pun intended!). How are you fixing and embedding the specimens, and what resin are you using? Also, what about the immunolabeling procedure? Are you using Protein A-gold or a "bridge antibody" like maybe goat anti-rabbit IgG and then streptavidin-gold?
Sorry I have more questions than answers. Maybe there's someone else out there that does this kind of work on a regular basis and can give you a more direct answer. If not, maybe we can help if you provide some additional details.
Best wishes, Bob **************************************** Robert (Bob) Chiovetti Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives / Systems Integrators / Analog & Digital Imaging *****************************************
It seems to me that the last time this "thread" on NSF document 122 and outside service work went through the listserv the issue of when NSF ownership of a piece of equipment ends came up......anyone recall something along these lines? If one uses the standard of 10% depreciation/yr, then any piece of equipment housed in a University - even though NSF-bought - has minimal value at the end of 10 years. Add on increasing maintenance and service costs as the equipment ages and you can be sure that there is a "crossover point" at which the University has more invested on an annual basis than the equipment is worth. At that point, ownership comes into question.
I appreciate that this too is a "fine point", but there is ownership, and there is ownership.
Winton
PS: I have no NSF-bought equipment
Dr. Winton Cornell Senior Research Associate & Supervisor, Microanalysis Laboratory Department of Geosciences The University of Tulsa 600 South College Tulsa, OK 74104-3189
The Electron Microscope Unit at the University of New South Wales has available for sale one JEOL WDS Spectrometer (in excellent condition - hardly used) suitable for the JEOL 840 / JXA- 8600S series of electron microscopes. Two light element crystals - STE (sterate) and TAP - thallium acid phthalate - are also available for sale. Also on offer is one set of an early 1980's vintage counting electronics/hv supply in a JEOL cabinet (~48" x 24" x 24"). The EM UNIT is seeking expressions of interest on any one or all of the above items. Please respond to me by Email if you are interested. Thankyou Barry EM UNIT UNSW
For Ren-Jye and others interested in preparing in particular: stained sections of polymers for TEM, and in general: all kinds of materials ranging from composite to catalysts;
may I recommend the following review:
H.K.Plummer - "Reflections on the use of microtomy for materials science preparation"
Microsc. Microanal. 3, 239-260 (1997)
Which draws on years of experience at the Ford Research Laboratory.
We ourselves are etchers, and only occasionally stainers.
+------------------------------------------------------------------------+ | 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 | +------------------------------------------------------------------------+
HELP US! We are interessed in the acquisition of a "second hand" (old fashioned)
SEM and TEM electronic microscopes. Free of charge is also accepted and appreciated. If you think you might help us in this matter, non hesitate give me a call:
Oncological Institute BUCHAREST-ROMANIA Corneliu Mateescu Ph.D cmateescu-at-mail.iob.ro =====================================================
I think it is important to consider the customer in this debate. We are in a slightly different situation here but have been involved in supplying a commercial service for many years. When we started providing this service we first calculated the true cost of the service. This cost incorporated all costs such as equipment price, depreciation, servicing, and operator time. Once this true cost is charged I think competition with outside services is then acceptable. Universities can provide a different element to a service which an outside service with a narrow focus cannot supply. This may be knowledge/expertise or new techniques. Is it right to deprive customers of these benefits simply because we are a University ? We have an excellent relationship with competing commercial services because we are not seen to exploit our position to undercut them. In fact our closest competitor ( six miles ) in the outside world sends work into us when they are under pressure. I should point out too that the last SEM bought for us was 17 years ago and we have funded all purchases of equipment from earnings since then. I know that our customers do not come to us because of price but because we provide a necessary service to them. Once a true commercial rate is charged I feel that the playing field is level and Universities should not be prevented from providing this service.
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: Garber, Charles A. {cgarber-at-2spi.com} To: MICROSCOPY BB {Microscopy-at-sparc5.microscopy.com}
We are looking to purchase a used EPMA, preferably a JEOL 733 or later.
Anybody got one languishing in the spare room?
Ritchie
Ritchie Sims Phone : 64 9 3737599 ext 7713 Department of Geology Fax : 64 9 3737435 The University of Auckland email : r.sims-at-auckland.ac.nz Private Bag 92019 Auckland New Zealand
The responses we received for the FREE SEM were overwhelming. In fact, there was a gentleman knocking at the door Monday morning and he is the lucky winner. If there is a change in plans, I will revert back to the list of inquirees and pick another winner.
} Hello! } } I am a TV researcher based in Vancouver, working on developing a TV } series about new initiatives in Microscopy. I am looking to find the } institution (s) and/or individual (s) doing the newest intiatives in the areas of: } } *Nanotechnology; } *Microscopy: techniques, technologies, Microvision, etc. } *Microelectronics; } *Microbiology: Digital Centre for Microbial Ecology, primordial } microbes? microbes on Mars, DNA microarrays, microbia physiology, } American Type Culture Collectin, microshells - shells less than 5 mm in size and microfossils; } *Microoptics } *Minerology: micrographs of rocks, crystals, etc. } *Micrometeorology } *Micromechanics: micromaching - Sensors, Actuators } *Microcirculation } } Please let me be more specific. Some possible topics, drawn from a wide range of scientific disciplines, might include: microscopy - technology, human parasites, insect flight, insect smells, nanotechnology, optical computing, fusion power, biological computers, targetted drug delivery, ocean microorganisms, the soup of life, cell regeneration, etc. } } I would appreciate any information. Since I'm looking for the newest, } not necessarily documented, initiatives in these areas - it is a little } difficult. } } PLEASE EMAIL ME DIRECTLY AT callain-at-intouch.bc.ca
I will be signing off soon - politics and early retirement strike again! Clearing my office! I want to thank you for your presence in my life in recent years. Also for responding to my questions - I really appreciate it. You are a great bunch of people. There is nothing like helpful support when you need it! I may be back in another guise. In the meantime, thanks, farewell and may the electrons be with you!
Keith Ryan Plymouth Marine Lab., UK PS - Daniele - don't cry! The e-mail address will still work, I hope. We'll meet again (now everybody wonders, who is this Daniele?!) XXX
Can you please send one to us. I have spend some time helping people with deciding on what they need. But this did not take off as I would have wished for. A opinion from a independent individual will help.
} One can make up a "decision tree" to help with this. If people are } interested, I can write up an article that walks through these steps to help } one decide on the most appropriate technology based on a set of needs. } -Ted Inoue } 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
Dear all please pass this to any relevant people. Please note imminent closing date.
RESEARCH ASSOCIATE/RESEARCH FELLOW (REF: A78/98)
CENTRE FOR MICROSCOPY AND MICROANALYSIS
The Centre for Microscopy and Microanalysis is the premier Microscopy Centre in Western Australia. In mid 1998 it was awarded a Western Australian Government Centre of Excellence in association with Curtin, Murdoch and Edith Cowan Universities.
The appointee will be required to carry out research in advanced microscopy. It is expected that the appointee will also take a leading role in the development of confocal microscopy, training and support of users, development of specimen preparation techniques, development of associated computer facilities and the support of research and teaching involving this instrument within the Centre. The minimum qualification required is a PhD in Science, Medicine or Agriculture together with experience in light and electron microscopy. Preference will be given to applicants with a substantial knowledge and skill in confocal microscopy and computing. The successful applicant must have strong interpersonal skills and the ability to work as part of the highly motivated group of academics at the Centre. The position is tenurable. For further information and copies of the selection criteria please contact either Associate Professor John Kuo on telephone (618 - international) OR (08) 9380 2765 or email jjskuo-at-cyllene.uwa.edu.au or Dr Gregory Pooley on telephone (618 - international) OR (08) 9380 2261 or email gdp-at-cyllene.uwa.edu.au or access the web link below.
SALARY RANGE: Level A $32,585 - $44,221 p.a. (Minimum starting salary for appointee with PhD will be $41,196 pa) Level B $47,946 - $56,937 p.a.
CLOSING DATE: 4 December 1998
Conditions of appointment will be specified in any offer of appointment which may be made as a result of this advertisement.
Written applications quoting reference number, telephone number, qualifications and experience and the names, addresses (including Email) and fax/telephone numbers of 3 referees should reach the Director, Human Resources, The University of Western Australia, Nedlands WA 6907, by the closing date.
http://jobs.uwa.edu.au/
The University is an equal opportunity employer and promotes a smoke-free work environment.
Brendan J. Griffin Centre for Microscopy and Microanalysis The University of Western Australia Nedlands, WA, AUSTRALIA 6907 ph 61-8-9380-2739 fax 61-8-9380-1087
Sorry that I didn't provide all the details on the original post, here they are:
We are attempting to label g-protein in Arabidopsis guard cells. Westerns indicate the protein is water soluble, but also associates with membranes, though it is not an integral membrane protein. Conventional fixation of the cells, with aldehydes, osmium, embedded in Spurrs yeilds good ultrastructure, but no labelling. For labelling we are fixing with 4% paraformaldehyde, 0.5% glutaraldehyde in 0.1 M phosphate buffer; dehydration in ethanol, embedding in LR White. The primary antibody is polyclonal, affinity purified IgG, the secondary is goat anti-rabbit conjugated to 10 nm colloidal gold. We are getting acceptable ultrastructure and labelling in E. coli (expressing the protein) processed this way, but not even a hint of labelling in plants.
We do have the resources to do cryo-work, but the equipment is all brand new, and we are still working out the details. Any suggestions on improving this protocol will be greatly appreciated. Thanks
#################################################### Michelle Peiffer Electron Microscope Facility for the Life Sciences The Biotechnology Institute for Research and Education 1 South Frear Lab University Park, PA 16802 814-865-0212 email:mlk101-at-psu.edu ####################################################
We have a Nikon TMD inverted epifluorescence microscope and a Nikon Labophot upright scope. We do not have an oil immersion lens on the inverted. Does anyone know if it would be a bad idea to take the oil immersion lens (Nikon Plan 100 (serial number 190370)) off of the upright scope and use it on the inverted? Can an oil immersion lens for an upright scope be used in the inverted position without harm?
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
Listers, A grad student in my TEM lab is requesting information regarding "calcium mapping." Her research involves calcium signalling during the metamorphosis of certain hydroids. She's using potassium pyroantimonate to precipitate free calcium during fixation/embedding. Precitate has been found in "unusual" places and calcium mapping has been suggested to verify that it is indeed calcium. Since "graduate student" is synonymous with "poverty" (her words) is there anyone out there who would offer free advice or low-cost mapping services. Please o' please (her words). Any response would be greatly appreciated. Contact can be made through me. Danke.
~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Winston W Wiggins, Supervisor 11/11/98 11:12:34 AM CRC-Electron Microscopy Lab. Ofc:704/355-1267 Carolinas Medical Center Fax:704/355-7648 P.O. Box 32861 Lab:704/355-7220 Charlotte,NC 28232-2861 USA Eml:wwiggins-at-carolinas.org ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
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I am using glycol methacrylate embedding and immunostaining to investigate polarity in small aggregates (100 microns) of hepatocytes. I have embedded the aggregates along with liver tissue and intestinal tissue (as controls) in JB4 glycol methacrylate following manufacturer's instructions. All attempts at staining so far have yielded results in the liver tissue only. No staining has been observed in the aggregates or the intestine which are significantly smaller in volume. I have been using trypsinization and several forms of "etching" I found in the literature (NaOH in 50% Ethanol/water, acetone). I am using mouse primary antibodies and goat-anti-mouse Oregon Green 488 as a secondary.
Any suggestions for getting results in the smaller tissues?
Thanks, Susan A. Fugett
Department of Chemical Engineering and Materials Science Phone: 612-625-8803 University of Minnesota 612-625-0808 421 Washington Ave SE Fax: 612-626-7246 Minneapolis, MN 55455 Email: fugett-at-cems.umn.edu
In a message dated 98-11-11 11:22:36 EST, wise-at-vaxa.cis.uwosh.edu-at-sparc5.microscopy.com writes:
{ { Does anyone know if it would be a bad idea to take the oil immersion lens (Nikon Plan 100 (serial number 190370)) off of the upright scope and use it on the inverted? Can an oil immersion lens for an upright scope be used in the inverted position without harm? } } Hi Bob,
You can certainly do this, it won't hurt a thing, in fact one of my customers has set up just such a situation on a Nikon inverted scope. But keep a couple of things in mind:
1. Use the bare minimum of oil that is necessary to fill the gap. Oil will tend to run down the lens, and this may cause a bit of a mess. Have lots of lens paper handy!
2. The oil lens probably has a fairly short working distance. This means that you may not be able to focus through a microscope slide or a culture plate, for example. But there are some options here, as well:
2A. If your specimen has to be on a microscope slide, you could seal the coverslip with paraffin wax or Vaseline and invert the whole slide, I suppose.
2B. If the specimen has to remain upright, you could place it on a thin coverslip and make a holder for the stage with an open space for the coverslip. There should be enough working distance in the lens to focus on the far (upper) surface of the coverslip.
3. If the specimen is still out of the focal range of the lens, you may have to modify the stage or make a simple flat metal plate that would substitute for the stage on the scope. My customer had to take this route for his setup, but it works just fine.
Hope this helps.
Cheers, Bob **************************************** Robert (Bob) Chiovetti Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives Systems Integrators Analog & Digital Imaging Systems *****************************************
Bob: We use our inverted Nikon with oil immersion lenses every day for the last 5 years with no trouble. Use a minimum amount of oil, clean well when you are done. There have been reports on the listserver (either microscopy or confocal) where users have gotten oil into the innards of the objective and some users have made dams out of various things (e.g., rubber o-rings) but we have avoided that. A NA 1.4 condenser on an upright scope is, of course, meant to be oiled in the same way and they generally survive (tho an MD ruined the condenser in one of my old labs by not cleaning up after himself when he was done). Good luck. Tom
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
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)
In a message dated 98-11-11 11:22:36 EST, wise-at-vaxa.cis.uwosh.edu-at-sparc5.microscopy.com writes:
{ { Can an oil immersion lens for an upright scope be used in the inverted position without harm? } } Bob,
One thing I forgot to mention: Several scope manufacturers (I don't know about Nikon) use a standard lens thread on their upright scopes and an "RMS" thread on their inverted scopes. If this is the case you will need an adapter collar which screws on the lens to convert it to an RMS thread for the inverted scope. I know Leica has such collars, so surely Nikon also has them if they are needed.
Keep in mind that using a collar on the lens *may* cause a slight parfocality problem.
Try gently screwing the lens into the inverted scope's nosepiece turret. If the lens does not mate properly, you will need an adapter collar.
Bob **************************************** Robert (Bob) Chiovetti Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives Systems Integrators Analog & Digital Imaging *****************************************
Thanks all - 23 responses to this posting (so far). I will summarize/respond in turn ASAP.
-----Original Message----- } From: Lehman, Ann Sent: Monday, November 09, 1998 10:37 AM To: 'MSA Listserver'
Winston Wiggins wrote: } } Listers, } A grad student in my TEM lab is requesting information } regarding "calcium mapping." Her research involves calcium signalling } during the metamorphosis of certain hydroids. She's using potassium } pyroantimonate to precipitate free calcium during fixation/embedding. } Precitate has been found in "unusual" places and calcium mapping has } been suggested to verify that it is indeed calcium.
What concentration of Ca is she looking for? The usual con- centrations involved in signalling are lower than micromolar, and are exceedingly difficult to impossible to determine with microanalytical techniques. The Somlyos have succeeded in seeing very low Ca concen- trations with EELS, and Marie Cantino reported using x-ray microanaly- sis to determine Ca (Proc MSA/MAS 1998) also at low levels. WDS could conceivably have sufficient sensitivity, but I don't know whether any- one has used it for these kinds of studies.
} Since "graduate student" is synonymous with "poverty" (her words) } is there anyone out there who would offer free advice or low-cost mapping } services. Please o' please (her words). Any response would be greatly } appreciated. Contact can be made through me.
Our facility has EDS (which can detect millimolar [Ca] or so, depending on the matrix), and we are a biotechnological resource, so we would be free (except for travel, hoterl, meals, etc.), but we are not likely to be suitable unless [Ca] is fairly high. Good luck. Yours, Bill Tivol
One of the vacuum pumps servicing our electron microprobe just gave up the ghost. We are faced with replacing the pump as this particlur pump from Edwards (an EDM-12) is obsolete - thus, rebuild kits are unavailable from Edwards. (if rebuilding were the way we wanted to go)
Edwards has a stock of a rebuilt pump that is a successor to ours (2 generations removed), with these rebuilt to factory specs. (this pump is the RV-12). I can get one for a reasonable price, which is about 60% of the cost of a new pump.
My questions to you:
1. how do you feel about the "vacuum quality" of rebuilt pumps vs. new pumps
2. how close do I have to get in pump specs?....the new pump has a capacity of 17.0 m^3/hr, while the old has (had) a capacity of 17.5 m^3/hr, i.e., there is about a 3% difference between them......should I shoot for a higher pumping capacity relative to the the old pumps?
Thanks, in advance, for your responses.
Winton
Dr. Winton Cornell Senior Research Associate & Supervisor, Microanalysis Laboratory Department of Geosciences The University of Tulsa 600 South College Tulsa, OK 74104-3189
So a little while ago I asked about prep'ing suspension cells for SEM. I got a lot of helpful suggestions which I will summarize & post here when I get the time. It ends up being that the guy grew them on gelatin coated coverslips and chamber slides which are WAY too big for my cpd. So I'm either going to have to try to cut or break them to make them fit. Or I was considering HMDS, has anybody out there used HMDS for things as delicate as cells? If you have could you e-mail me with protocols? I have the one that works for bugs and I have the HMDS, I've just never used it. Any help you send my way will be greatly appreciated. I'm getting sooo smart from y'all's ideas that my head hurts.
Going quietly into the SEM room,
Paula :-)
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
Try etching for 3-5 minutes in sodium ethoxide soln 1:1 with toluene, = then go to 100% EtOH down to water, buffer, etc. Ethoxide Solution: 75gm NaOH in absolute ethanol; let stand 10-14 days or until pellets = dissolve with occasional stirring. Solution should turn brown when = ready. This works well with tissue embedded in Araldite 502 (and may = work with JB4 media) allowing all types of histochemistry procedures and = immunohistochemistry.
Hank Adams Cell Biology Integrated Microscopy Core Baylor College of Medicine One Baylor Plaza Houston, Tx 77030
-----Original Message----- } From: Susan Fugett [SMTP:fugett-at-cems.umn.edu] Sent: Wednesday, November 11, 1998 5:26 PM To: microscopy listserver
I am using glycol methacrylate embedding and immunostaining to=20 investigate polarity in small aggregates (100 microns) of hepatocytes. I =
have embedded the aggregates along with liver tissue and intestinal=20 tissue (as controls) in JB4 glycol methacrylate following manufacturer's =
instructions. All attempts at staining so far have yielded results in = the=20 liver tissue only. No staining has been observed in the aggregates or = the=20 intestine which are significantly smaller in volume. I have been using=20 trypsinization and several forms of "etching" I found in the literature=20 (NaOH in 50% Ethanol/water, acetone). I am using mouse primary = antibodies=20 and goat-anti-mouse Oregon Green 488 as a secondary.
Any suggestions for getting results in the smaller tissues?
Thanks, Susan A. Fugett
Department of Chemical Engineering and Materials Science Phone: 612-625-8803 University of Minnesota 612-625-0808 421 Washington Ave SE Fax: 612-626-7246 Minneapolis, MN 55455 Email: fugett-at-cems.umn.edu
This is a multi-part message in MIME format. --------------0ACCEBDC64F1D9ABE2ADF72B Content-Type: text/plain; charset=us-ascii Content-Transfer-Encoding: 7bit
"Vacuum quality" which really means base pressure and backstreaming is more effected by the choice of pump oil than mechanical matters. If the rebuild was done properly there is no problem.
Pumping speed is just that. The new pump will be 3% slower to pump down. The length of vacuum hoses and the presence of foreline traps effects pumping speed more. The only slow pumping speed problem I have seen was where a new microscope was installed with a trap and too long of a vacuum line so that the evacuation control electonics and valves went into oscillation at crossover because of slow pumping and some outgassing.
Ronald Vane XEI Scientific 650-369-0133
Winton Cornell wrote:
} Folks: } } One of the vacuum pumps servicing our electron microprobe just gave up the } ghost. We are faced with replacing the pump as this particlur pump from } Edwards (an EDM-12) is obsolete - thus, rebuild kits are unavailable from } Edwards. (if rebuilding were the way we wanted to go) } } Edwards has a stock of a rebuilt pump that is a successor to ours (2 } generations removed), with these rebuilt to factory specs. (this pump is } the RV-12). I can get one for a reasonable price, which is about 60% of the } cost of a new pump. } } My questions to you: } } 1. how do you feel about the "vacuum quality" of rebuilt pumps vs. new pumps } } 2. how close do I have to get in pump specs?....the new pump has a capacity } of 17.0 m^3/hr, while the old has (had) a capacity of 17.5 m^3/hr, i.e., } there is about a 3% difference between them......should I shoot for a } higher pumping capacity relative to the the old pumps? } } Thanks, in advance, for your responses. } } Winton } } Dr. Winton Cornell } Senior Research Associate & Supervisor, Microanalysis Laboratory } Department of Geosciences } The University of Tulsa } 600 South College } Tulsa, OK 74104-3189 } } phone: 918-631-3248 } fax: 918-631-2091 } e-mail: wcornell-at-centum.utulsa.edu
} Or I was considering HMDS, has anybody out there used HMDS for } things as delicate as cells?
Shirley Pinchuck, in this lab, has done a fair amount of work using HMDS, including comparisons of HMDS with other methods such as CPD, cryo-SEM, etc. I will ask her to fax you her protocols as well as an abstract of a conference presentation on some of the comparative work.
I hope this helps.
Regards
Robin
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
There should not be any difference in performance between a new pump and a factory rebuilt one - they should both acheive the original vacuum specification. Obviously the rebuilt one may not look as shiny but we have been happy with rebuilt pumps over the years.
I assume that the pump is backing a high vacuum pump of some sort in which case it has to meet the specification for that, 3% is not significant. If your microprobe uses a Diffstack 160 (guessing at 1 vacuum supplier) then the RV12 is the currently reccomended backing pump.
Regards, Ron
} Folks: } } One of the vacuum pumps servicing our electron microprobe just gave up the } ghost. We are faced with replacing the pump as this particlur pump from } Edwards (an EDM-12) is obsolete - thus, rebuild kits are unavailable from } Edwards. (if rebuilding were the way we wanted to go) } } Edwards has a stock of a rebuilt pump that is a successor to ours (2 } generations removed), with these rebuilt to factory specs. (this pump is } the RV-12). I can get one for a reasonable price, which is about 60% of the } cost of a new pump. } } My questions to you: } } 1. how do you feel about the "vacuum quality" of rebuilt pumps vs. new pumps } } 2. how close do I have to get in pump specs?....the new pump has a capacity } of 17.0 m^3/hr, while the old has (had) a capacity of 17.5 m^3/hr, i.e., } there is about a 3% difference between them......should I shoot for a } higher pumping capacity relative to the the old pumps? } } Thanks, in advance, for your responses. } } Winton } } } Dr. Winton Cornell } Senior Research Associate & Supervisor, Microanalysis Laboratory } Department of Geosciences } The University of Tulsa } 600 South College } Tulsa, OK 74104-3189 } } phone: 918-631-3248 } fax: 918-631-2091 } e-mail: wcornell-at-centum.utulsa.edu } } } }
=========================================================================== 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. ============================================================================
Our group currently has a JEOL FX 2000 TEM available for sale to anyone interested. The system includes a cold and hot stage, PC driven X-ray system,and STEM attachment. Please respond if interested.
M.W. Rigler, Ph.D. MAS, Inc. Suwanee GA 770-866-3218
Hi group, I am posting this message to resume the problem that was posted some time ago (17.09.98 ' latex microsheres') with request on info about particle suppliers. I would like to thank everybody who replied but the problem was that NO ONE company from the list we received had right particles(1-10 nm, optically transparent at a given wavelength region and absorbing at different given region). Finally I have found the best source for not only me as I guess : SPHEROTECH INC, 847-680-8922, http://www.spherotech.com with the widest range of particles for microscopy needs. Best regards Dmitri Lapotko Luikov Heat and Mass Transfer Insitute Minsk Belarus
Bob, Having been a dealer rep for Nikon, I respond this way. You can invert the lens and use it without adapters or problems with parfocality, however, the working distance is very limited and would only work when the sample is resolved through a cover glass. This tells you that the sample could just as easily be viewed on a compound microscope. Nikon does offer long working distance lenses specifically designed for the TMD. I do not recall a 100X oil ever being offered, but they do offer a 60X ELWD dry.
I am teaching histology and microtechnique to advanced high school students. I need a source of good prepared demonstration slides. Somewhere along the way I lost my collection which I made when I worked in a medical lab. Because I am paying for the slides myself they need to be economical. I bought some from Edmund Scientific and found that they were unuseable, pathetic junk. Can anyone suggest good quality sources for prepared slides that I might be able to afford. I am especially interested in series of slides of the same specimen that use a variety of stains. I would also be interested in purchasing slides from any of the experts on this mailing list who must have great examples of their skills which would be appropriate for educational purposes.
It is possible that the statement below about the speed of the pump being 3% slower may not be relevant to your situation. What should be taken into consideration in selecting a pump is the effective pump speed. The effective pump speed is the speed at the vacuum chamber port when the capacitance of the tubing from the pump to the vacuum system is taken account. It works like parallel resistors. The effective pump speed is given as
where S(pump) is the pump speed and C(tubing) is the capacitance of the tubing. A system can easily become capacitance limited if C(tubing) { { S(pump). When this happens, Seff=S(pump). If this is the case, the 3% difference in pump speeds will not matter. Standard books on vacuum science can give you the values of capacitance for the length and diameter of tubing, elbows, valves, and how they are added to give and effective value to the pump.
So, the moral of this story is that you should always consider your total vacuum system when selecting a pump.
-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: Ronald Vane To: Winton Cornell Cc: Microscopy-at-Sparc5.Microscopy.Com -----------------------------------------------------------------------.
Winton and all:
The rebuilt pump should be just fine. Go for it.
"Vacuum quality" which really means base pressure and backstreaming is more effected by the choice of pump oil than mechanical matters. If the rebuild was done properly there is no problem.
Pumping speed is just that. The new pump will be 3% slower to pump down. The length of vacuum hoses and the presence of foreline traps effects pumping speed more. The only slow pumping speed problem I have seen was where a new microscope was installed with a trap and too long of a vacuum line so that the evacuation control electonics and valves went into oscillation at crossover because of slow pumping and some outgassing.
Ronald Vane XEI Scientific 650-369-0133
Winton Cornell wrote:
} Folks: } } One of the vacuum pumps servicing our electron microprobe just gave up the } ghost. We are faced with replacing the pump as this particlur pump from } Edwards (an EDM-12) is obsolete - thus, rebuild kits are unavailable from } Edwards. (if rebuilding were the way we wanted to go) } } Edwards has a stock of a rebuilt pump that is a successor to ours (2 } generations removed), with these rebuilt to factory specs. (this pump is } the RV-12). I can get one for a reasonable price, which is about 60% of the } cost of a new pump. } } My questions to you: } } 1. how do you feel about the "vacuum quality" of rebuilt pumps vs. new pumps } } 2. how close do I have to get in pump specs?....the new pump has a capacity } of 17.0 m^3/hr, while the old has (had) a capacity of 17.5 m^3/hr, i.e., } there is about a 3% difference between them......should I shoot for a } higher pumping capacity relative to the the old pumps? } } Thanks, in advance, for your responses. } } Winton } } Dr. Winton Cornell } Senior Research Associate & Supervisor, Microanalysis Laboratory } Department of Geosciences } The University of Tulsa } 600 South College } Tulsa, OK 74104-3189 } } phone: 918-631-3248 } fax: 918-631-2091 } e-mail: wcornell-at-centum.utulsa.edu
HMDS can work fine for cells. You might have to play a bit to find the best transition series, but I'd start with 3:1-} 1:1-} 1:3 absEtOH:HMDS. Drying is the bigger question: how fast at what temperature? If I may, we ran an article on this in the May '97 issue of Microscopy Today. If you don't have a copy, I can see if we've got an extra one.
Information on HMDS has also been posted on the U Florida Tips & Tricks of Microscopy pages: http://www.biotech.ufl.edu/~emcl/tips.html
I can send more details if you wish.
Phil
} It's me again, } } So a little while ago I asked about prep'ing suspension cells for } SEM. I got a lot of helpful suggestions which I will summarize & post here } when I get the time. It ends up being that the guy grew them on gelatin } coated coverslips and chamber slides which are WAY too big for my cpd. So } I'm either going to have to try to cut or break them to make them fit. } Or I was considering HMDS, has anybody out there used HMDS for } things as delicate as cells? If you have could you e-mail me with } protocols? I have the one that works for bugs and I have the HMDS, I've } just never used it. } Any help you send my way will be greatly appreciated. I'm getting } sooo smart from y'all's ideas that my head hurts. } } } Going quietly into the SEM room, } } } Paula :-) } } 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
****be famous! send in a tech tip or question*** Philip Oshel Technical Editor, Microscopy Today PO Box 620068 Middleton, WI 53562 (608) 833-2885 oshel-at-terracom.net
This is the promised follow up on my question (posted 11/6/98) about color CCD cameras, included is my original posting and the responses I received to date. I really appreciated the "low key" and informative responses of some of the vendors.
I should add that I found some of the information that helped me frame my questions from this site: http://www.soundvisioninc.com/howdcw.htm
Doug Cromey
--------------the original question---------------------
} Microscopy Listers, } } We are in the early stages of evaluating a digital imaging system here (for } light microscopy). I'm trying to understand the different ways that CCD } cameras can be or are used to acquire color images. There seem to be } several ways this is done: } } * Single chip Monochrome CCD array with some type of filter in } front of the camera to allow it to acquire sequential red, } green & blue images and then software to "reassemble" the } images into a color image. } * 3 Chip camera, with each chip assigned (filtered for) red, } green & blue and then software to "reassemble" the images into } a color image. } * Single chip CCD with some type of color mosaic "mask" on the } chip to acquire the red, green & blue parts of the image and } then software to "reassemble" the images into a color image. } * CCD array where the image is "scanned" either by moving optical } elements or moving the CCD array to acquire a high pixel count } with a fairly small sensor. These would be the slowest for } acquisition, but I gather they give a lot of "bang for the } buck (euro)". } * Others? } } What are the Pros & Cons of these different types of cameras? Is there a } WWW or published resource that could help me sort this out? } } Our interest here is primarily in acquiring static images (not real-time } video) from low light level fluorescence, DIC and/or bright field. We } would like the camera to be sensitive enough for quantitation if needed. } I'd also like to be able to advise others here who may have different } requirements. I'm not specifically "fishing" for sales pitches (I already } have plenty of glossy literature, I'm just trying to make sense out of it). } } I would be happy to take replys "off-list" and post a summary. } } Yours, } Doug Cromey
----------reply--------------
} From: RCHIOVETTI-at-aol.com
good for resolution and, if properly used, color fidelity but not good for motion or dynamic or time sensitive subjects (like fading flourescence)
} * 3 Chip camera, with each chip assigned (filtered for) red, } green & blue and then software to "reassemble" the images into } a color image.
reasonably good color fidelity but the dichroic filters suck up lots of the light and require special care with the selection of the adapter - dichroics what parallel light or you'll get funny color fringes and shifts
} * Single chip CCD with some type of color mosaic "mask" on the } chip to acquire the red, green & blue parts of the image and } then software to "reassemble" the images into a color image.
color fidelity is not as good - most have 1/2 green, 1/4 blue and 1/4 red but at least one (Panosonic GP-KS1000) has more pixels (900,000 vs 410,000) and a different color array of 1/3 each of R/G/B
} * CCD array where the image is "scanned" either by moving optical } elements or moving the CCD array to acquire a high pixel count } with a fairly small sensor. These would be the slowest for } acquisition, but I gather they give a lot of "bang for the } buck (euro)".
scanned arrays can give unbelivable resolution - I just saw a one this weeek that used a 10K Kodak linear array! but the time to scan is often the limiting factor with all that that implies (see - http://betterlight.com)
} * Others?
CMOS - new technology, less expensive (see http://www.soundvisioninc.com) Digital cameras - if you never need "video" these can be your salvation, just decide what you need to do and find one that does it (see http://www.electrim.com or the guy from DVC ) there are also CID cameras (low blooming)
There are also a host of all of the above with low light capabilities either through such technology back thining or on chip integration
} } What are the Pros & Cons of these different types of cameras? Is there a } WWW or published resource that could help me sort this out? } } Our interest here is primarily in acquiring static images (not real-time } video) from low light level fluorescence, DIC and/or bright field. We } would like the camera to be sensitive enough for quantitation if needed. } I'd also like to be able to advise others here who may have different } requirements. I'm not specifically "fishing" for sales pitches (I already } have plenty of glossy literature, I'm just trying to make sense out of it). } } I would be happy to take replys "off-list" and post a summary. } } Yours, } Doug } .................................................................... } : Douglas W. Cromey, M.S. Dept. of Cell Biology & Anatomy : } : Research Specialist, Principal University of Arizona : } : (office: AHSC 4212A) P.O. Box 245044 : } : (voice: 520-626-2824) Tucson, AZ 85724-5044 USA : } : (FAX: 520-626-2097) (email: doug-cromey-at-ns.arizona.edu): } :...................................................................: } http://www.pharmacy.arizona.edu/exp_path.html } Home of: "Microscopy and Imaging Resources on the WWW"
BTW - I don't sell any of this stuff.
---------------------reply-----------------------
} From: Thomas A Baginski {tombg-at-bictom.usuf1.usuhs.mil}
Dear Australians,
I had already deleted the message, when I found myself in contact with someone with experience in confocal microscopy who might be interested in the Confocal job. Could you please re-send the advert to me?
+------------------------------------------------------------------------+ | Robert H.Olley Phone: | | J.J.Thomson Physical Laboratory {direct line +44 (0) 118 9318572 | | University of Reading {University internal extension 7867 | | Whiteknights Fax +44 (0) 118 9750203 | | Reading RG6 6AF Email: R.H.Olley-at-reading.ac.uk | | England URL: http://www.reading.ac.uk/~spsolley | +------------------------------------------------------------------------+
I am working with a mixture of organic and water on observation using optical microscope. Here is no contrast between water and organic. I need to know if the "droplets" observed are water droplets, or organic. Is there a die or similar available to colour water so that there is a contrast between water and organic?
Zhaoxia Zhou Department of Materials Engineering Brunel University, UK Tel: +44-1895-274000-2968(o) E-mail: Zhaoxia.zhou-at-brunel.ac.uk
Could something be sent to this list too please - it would certainly interest me.
thanks
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD Tyne and Wear UK
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk ---------- } From: ROBIN CROSS To: Paula Sicurello Cc: microscopy
Hello Paula
} Or I was considering HMDS, has anybody out there used HMDS for } things as delicate as cells?
Shirley Pinchuck, in this lab, has done a fair amount of work using HMDS, including comparisons of HMDS with other methods such as CPD, cryo-SEM, etc. I will ask her to fax you her protocols as well as an abstract of a conference presentation on some of the comparative work.
I hope this helps.
Regards
Robin
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
by newton.wadsworth.org (8.8.8/8.8.8) with SMTP id NAA29824; Thu, 12 Nov 1998 13:06:42 -0500 (EST) Sender: tivol-at-wadsworth.org Message-ID: {364B210D.41C6-at-wadsworth.org}
Zhaoxia Zhou wrote: } } I am working with a mixture of organic and water on observation using } optical microscope. Here is no contrast between water and organic. I need } to know if the "droplets" observed are water droplets, or organic. Is there } a die or similar available to colour water so that there is a contrast } between water and organic? } Dear Zhaoxia, Colored ions could solve your problem (depending on whether there is significant partitioning into the organic). There is also a potential problem, since the ions might interfere with whatever else is happening. Cu++ is a nice blue color, and the intensity is greatly enhanced in the presence of NH4+. If you can add CuSO4 and NH4OH to the water without any problem, then you should see contrtast. However, if the organic has the possibility of forming N ligands, e.g., phenanthroline, the Cu could also color the organic. Also, if the organic has COOH groups with high pK's then the NH4OH could cause mixing of the organic and H2O due to the COO- groups formed at high pH. Good luck. Yours, Bill Tivol
I was wanting to know if anyone has used any wax or etch resist which will protect against warm KOH. I need to mask a Si/SiO2/Si wafer (Silicon on Insulator) to preferentially remove a the thick Si backside of my specimen and leave the surface Si/SiO2 bilayer intact for TEM imaging. I was going to use warm KOH (50 C) to remove the Si but I need to mask the surface and define a window on the backside Si to etch through (the KOH will stop at the SiO2 layer). Any suggestions?
John --------------------------------------- John Michael Glasko Materials Science and Engineering Dept. 2142 Burlington Labs/Yarborough St Raleigh, NC 27695-7916 USA tel: (919)-515-7217, fax: (919)-513-1699 jmglasko-at-unity.ncsu.edu
I've never prepared a Schiff reagent before and I'm having problems. I've been using the protocol in Presnell and Schreibman's edition of Humason's Animal Tissue Techniques, which recommends: *** basic fuchsin 0.5-1.0 g water 85 ml sodium metabisulfite 1.9 g 1.0 N HCl 15 ml
Shake for 2 hr or let sit overnight. Add 2 g of activated carbon, shake for about a minute, then filter. The solution should be water-clear. ***
Well, my solution is not water clear. In my hands, the dye never dissolves completely, and the activated carbon has absolutely no effect. P/S say that if the solution doesn't clear the charcoal is old, but I've been using a fresh batch. The resulting filtered solution is pale orange.
Any suggestions?
Gary Radice 804-289-8107 Department of Biology 804-289-8233 (FAX) University of Richmond gradice-at-richmond.edu Richmond VA 23173
Engineered Carbons, Inc., a subsidiary of Ameripol Synpol Corporation, is replacing its Quantimet 720 with digital imaging instrumentation. Is this equipment of value to anyone in the microscopy community? Included with the equipment is a PDP11E/05 tape recorder, a TV video disk recorder, a DECwriter, and other associated parts.
The interested individual/institution would be responsible for transport from Borger, TX (near Amarillo).
Chuck Butterick Engineered Carbons, Inc. P.O.Box 2381 1111 Penn Avenue Borger, TX 79008
I have been asked to find out if anyone can share any information about silicon nitride or silicon oxide supports for TEM. We are interested in 100 nm thick plates or sheets - where to get them or how to make them; and on my part, how they are used.
Thanks in advance,
Pat Hales McGill University Dept. of Anatomy & Cell Biology hales-at-med.mcgill.ca
Gary, that is not an uncommon problem. It depends on your source of = basic fucshin which is if my memory serves me right is a combination of = pararosaline dyes. Some sources are better than others and I went thru = quite a few. I had absolutely perfect results with some produced from = Sigma under pararosaniline in their catalogue, though I don't remember = which salt I used. The solution came out perfectly clear and I stored = aliquots at minus 20C and it seemed quite stable. =20
Hank Adams Cell Biology Integrated Microscopy Core Baylor College of Medicine One Baylor Plaza Houston, Tx 77030
Gary, It has been awhile since I worked with the Schiff reagent but I used to mix up large batches of it. Firstly, the formula that I have (Lillie (1954) is slightly different:
basic fuchsin (C.C.) 1 gm (best to use a dye certified by the Biological Stain Commission) distilled water 80 ml NaHSO3 is 2 gm or 1.9 gm Na2S2O5 N HCl 20 ml
I noticed that this formula uses more bisulfite and more HCl -- both of which generate more SO2 for decolorization of the fuchsin.
This should go in a tightly stoppered container (to keep in the SO2 which does the decolorization of the fuchsin) and shaken every 10 minutes for a period of 2 hr. Add 500 mg finely powdered fresh charcoal (we used coconut charcoal). Charcoal must be finely ground (like talc). You can do this twice to remove any residual color. Sometimes I even added charcoal, shake and store in a refrigerator overnight. Filter through several layers of paper filter media. When colorless store at 5 deg C. If white precipitate shows up, it can be refiltered. Pink color means time to discard it.
Since your stain is not dissolving, something may be wrong with the fuchsin. Lack of destaining may be caused by not enough bisulfite or loose fitting container. Most often, we did not even need to use the charcoal.
} I've never prepared a Schiff reagent before and I'm having problems. I've } been using the protocol in Presnell and Schreibman's edition of Humason's } Animal Tissue Techniques, which recommends: } *** } basic fuchsin 0.5-1.0 g } water 85 ml } sodium metabisulfite 1.9 g } 1.0 N HCl 15 ml } } Shake for 2 hr or let sit overnight. Add 2 g of activated carbon, shake for } about a minute, then filter. The solution should be water-clear. } *** } } Well, my solution is not water clear. In my hands, the dye never dissolves } completely, and the activated carbon has absolutely no effect. P/S say that } if the solution doesn't clear the charcoal is old, but I've been using a } fresh batch. The resulting filtered solution is pale orange. } } Any suggestions? } } Gary Radice 804-289-8107 } Department of Biology 804-289-8233 (FAX) } University of Richmond gradice-at-richmond.edu } Richmond VA 23173
#################################################################### 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 ####################################################################
It's been a while since I made my own Schiff's but I remember the frustration of the filtrate coming out colored (pink or orange) even after repeated shaking with activated charcoal. The protocol that I used was different from the one you describe (but close I think) so I hope this helps. The problem was that the flasks I was filtering into weren't completely dry, i.e. I would filter into them, rinse them and then filter into them again thinking "this is OK because the Schiff's solution is mostly water anyway." Water is an aldehyde and reacts with Schiff's to form a pink color. By ensuring that glassware is completely dry you should achieve a colorless Schiff's solution after a couple of charcoal treatments. Water in the atmosphere may be responsible for the sneaky way Schiff reagent has of turning from an insignificant appearing clear spill into an angry red stain on hands, clothes and benchtops. This is a good indicator of how sloppy one is being in the lab. Get the stuff on your hands a couple of times and you'll learn to be careful. BTW, this sort of trouble with home-brew Schiff's has inspired many people to buy the stuff ready made. Good luck, John } ----------------------------------------------------------- } } I've never prepared a Schiff reagent before and I'm having problems. I've } been using the protocol in Presnell and Schreibman's edition of Humason's } Animal Tissue Techniques, which recommends: } *** } basic fuchsin 0.5-1.0 g } water 85 ml } sodium metabisulfite 1.9 g } 1.0 N HCl 15 ml } } Shake for 2 hr or let sit overnight. Add 2 g of activated carbon, shake for } about a minute, then filter. The solution should be water-clear. } *** } } Well, my solution is not water clear. In my hands, the dye never dissolves } completely, and the activated carbon has absolutely no effect. P/S say that } if the solution doesn't clear the charcoal is old, but I've been using a } fresh batch. The resulting filtered solution is pale orange. } } Any suggestions? } } Gary Radice 804-289-8107 } Department of Biology 804-289-8233 (FAX) } University of Richmond gradice-at-richmond.edu } Richmond VA 23173
________________________ C. John Runions, Ph.D. Section of Ecology and Systematics Corson Hall Cornell University Ithaca, New York USA 14853
I have gotten some pretty good TEM's of a negative stained bacteriophage prep but, unlike some published photos of T4 and H. influenze phage, I don't get any fine details like striations in the tail. Essentially I get nice looking hexagons sitting on top of tails but no fine structure. I have tried staining with 1% PTA or 2% uranyl acetate or vandium. I have tried 2 basic protocols:
1. putting a drop of the prep on a carbon coated grid for 30 to 120 sec, wicking the drop off, rinsing with a drop of water, and then adding a drop of stain and almost immediately removing it. Skipping the rinse doesn't seem to help nor does lengthing the staining time.
2. putting a 5 ul drop of the prep on the grid and then adding a 5 ul drop of stain.
Are there any negative staining experts out there with hints for getting the best image?
TIA, tom
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility
3 Tucker Hall Division of Biological Sciences University of Missouri Columbia, MO 65211 (573)-882-4712 (voice) (573)-882-0123 (fax)
Pat Hales wrote: ================================================== I have been asked to find out if anyone can share any information about silicon nitride or silicon oxide supports for TEM. We are interested in 100 nm thick plates or sheets - where to get them or how to make them; and on my part, how they are used. ================================================= Most of the answers to the questions are found on our website URL http://www.2spi.com/catalog/instruments/silicon-nitride.html
The requested thickness is one of our standard thicknesses, and the short answer is that they are used just like a normal TEM grid since the outer support silicon is 3 mm diameter and can fit into any standard 3 mm grid holder. Depending on the type of work you are doing, you might want to consider something thinner than 100 um.
The properties of Si3N4 in this thickness are such that there is no such thing as a free standing "sheet" since at that thickness it would instantly curl up on itself into a roll.
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: www.2spi.com ############################ ==================================================
} Could something be sent to this list too please - it would certainly } interest me. } } thanks } } Malcolm Haswell
Malcolm,
This is the text of the article from the May '97 Microscopy Today. The U. =46lorida web site and Shirley Pinchuck mentioned in Robin Cross' post will have more information.
Phil
HMDS and Specimen Drying for SEM:
Hexamethyldisilizane (HMDS) is an excellent method of chemical drying of hydrated specimens. There are several variables involved in its use, the most easily controlled being the number of transitional steps from 100% ethanol (EtOH) to 100% HMDS and the drying temperature. Fixation and dehydration are the same for both HMDS and CPD. Once the specimen is in the final 100% ethanol, it must then be transferred to 100% HMDS through a graded series of ethanol-HMDS mixtures. This can follow one of four basic paths:
Ratio absEtOH : HMDS starting from 100% EtOH going to 100% HMDS
1)100% E =3D} 1:1=3D} 100% H 2)100% E =3D} 2:1=3D} 1: 2=3D} 100% H 3)100% E =3D} 2:1=3D} 1:1=3D} 1: 2=3D} 100% H 4)100% E =3D} 3 :1 =3D} 1:1=3D} 1: 3=3D} 100% H (Extra gradations may be added as needed, for instance between the 3 :1 - 1:1 and 1:1 - 1: 3 steps)
After the final transitional step, make 3 changes in HMDS (the last two steps can sometimes be skipped). Dry from the last 100% HMDS step, or exchange with new 100% HMDS one final time then dry. There should be just enough HMDS in the container to cover the specimen, any more just increases the drying time. Time in these steps will usually be the same as that used in the final 100% EtOH steps. However, the time can apparently be extended will little ill effect on the sample. Incomplete transition from EtOH to HMDS is a worse source of problems than extended time in HMDS. Choice of steps is basically determined by sample density, and the permeability (to HMDS and ethanol) of the least permeable structures in the sample. Microorganisms can usually be done with the first series, animal tissues need the second or fourth series, and plants require the fourth series, or even more gradations because of their cell walls.
Drying is done at either: 25=BA C =3D room temperature 8 hr =3D} overnight 37=BA C } 45=BA C }1=3D} 4hr 60=BA C } (Drying time by both temperature and volume of fluid.) The higher the temperature, the shorter the drying time, but the quality of results may also vary. Microscopic unicellular algae did best at 60=BA C, fish skin at room temperature, bacteria equally well at room temperature and 60=BA C. HMDS may have a significant time advantage over CPD. If more specimens are being processed that can fit in the CPD chamber, then the times in the transitional fluids and for drying will be much less than the time necessary for CPD. The greater the number of samples that can be batch processed, the greater the time advantage for HMDS. HMDS has another advantage: if you can find containers that seal tightly enough (HMDS likes to evaporate given any chance at all), samples can be collected in the field, processed to 100% HMDS, then stored in vials filled with HMDS and transported long distances from remote sites - like from Antarctica to Chicago, Illinois. The samples are protected by the fluid, and at least for some specimens, so fewer artifacts than specimens stored in fixatives or alcohol. Dried specimens are of course fragile. HMDS is not the cure-all for specimen drying. It can introduce it's own distorting artitfacts and some samples, biological ones in particular, still shrink after drying as they do with CPD or freeze-drying. Some specimens do poorly when dried from HMDS. Plant tissues in particular may not do well, and may be better off dried by CPD. Also, if the specimen is going to be studied for elements that are labile, or in solution, standard fixation and dehydration methods won=EDt work. Cryo techniques must be used, and if the specimen is to be examined in an unfrozen state, for example to look at structures and elements within cavities that would be obscured by ice if left hydrated, then the specimen must be dried by freeze-drying methods. All of this information is empirical. Theoretical explanations and any modifications for particular samples are welcome! A final note: HMDS must be used in a flume hood! A sniff of it will clear the sinuses back to the foramen magnum.
--MT
****be famous! send in a tech tip or question*** Philip Oshel Technical Editor, Microscopy Today PO Box 620068 Middleton, WI 53562 (608) 833-2885 oshel-at-terracom.net
Instead of etching the Si off to prepare a Plan View TEM sample, I suggest mechanically thinning the Si substrate. Allied High Tech is the manufacture of a tool, the MultiPrep, which is a semi-automatic tool for preparing SEM, TEM cross sections and TEM plan view samples in addition to its other capabilities.
I am an employee of Allied and have an interest providing you with this information. If you would like to receive more detailed information please contact me or visit our website http://www.alliedhightech.com
Sincerely,
Ed Hirsch
} Hi, } } I was wanting to know if anyone has used any wax or etch resist } which will protect against warm KOH. I need to mask a Si/SiO2/Si } wafer (Silicon on Insulator) to preferentially remove a the thick Si } backside of my specimen and leave the surface Si/SiO2 bilayer intact } for TEM imaging. I was going to use warm KOH (50 C) to remove the Si } but I need to mask the surface and define a window on the backside Si } to etch through (the KOH will stop at the SiO2 layer). Any } suggestions? } } John } --------------------------------------- } John Michael Glasko } Materials Science and Engineering Dept. } 2142 Burlington Labs/Yarborough St } Raleigh, NC 27695-7916 } USA } tel: (919)-515-7217, fax: (919)-513-1699 } jmglasko-at-unity.ncsu.edu } } ************************************************* Edward A. Hirsch Product Application Specialist Allied High Tech 2376 East Pacifica Place Rancho Dominguez, CA 90220 ph: (919) 846-9628 vm:(800)675-1118 x245 fx: (310)762-6808 http://www.alliedhightech.com *************************************************
I need information regarding use of low vacuum SEM in analysis of tissue culture cells combined with immunogold labelling. Has anyone tried? Most of the images I've seen from the manufacturer's brochures are of algae, insects or foods. I'd appreciate comments.
Corazon D. Bucana UT M.D. Anderson Cancer Center Houston, Texas
} 1. putting a drop of the prep on a carbon coated grid for 30 to 120 } sec, wicking the drop off, rinsing with a drop of water, and then adding a } drop of stain and almost immediately removing it. Skipping the rinse } doesn't seem to help nor does lengthing the staining time.
} 2. putting a 5 ul drop of the prep on the grid and then adding a 5 ul } drop of stain.
a) Did you 'glow discharge' the grid shortly before you adsorbed the drop? b) Have you tried Uranyl Formiate solution? (I think 0.75%w/w and pH around 4 should be good.) c) Have you tried lower concentration of Uranyl salt? Maybe the fine details are embedded in a thick salt layer so that you don't get any contrast there.
The procedure I normally use is: - prepare parafilm with at least 2 droplets of water (or sometimes buffer?) and another two of stain solution - glow discharge the grid for around 10 to 20 s (blue-white) - clamp it into tweezer - put 5um drop on the grid and wait... whatever.. 30 s, 60 s.... - touch with filter paper to soak solution away (blotting) - wash on first drop of water by touching the surface, blot again - repeat above step at least once - then go to stain droplet and do the same - the second stain droplet is then used to do the staining procedure, so touch the surface and wait for a few seconds (lets say 10 to 20) - blot a last time and that's it
You may not need all these steps, but I have been very successful with this technique and don't see any reason not to do it this way. If you don't have Uranyl Formiate, you can try the upper protocol with Uranyl Acetate (2% or lower). My experience is that usually it should work as nicely as UFo, but I was told that UAc does 'melt' more strongly and more quickly in the electron beam. I am looking forward to other recipies. I am very sure that you can find as many recipies as there are microscopists...
By the way, I have never been successful using PTA. Even with very sensitive proteins which don't like a low pH, I always had the best results in using Uranyl solutions.
Best regards,
Bettina
*** Bettina Wolpensinger Electron Microscope Unit University of New South Wales Sydney, NSW 2052, AUSTRALIA phone: +612 9385 6390 fax: +621 9385 6400 b.wolpensinger-at-unsw.edu.au http://srv.emunit.unsw.edu.au ***
Pale yellow colored Schiff-solutions are usable as long as they are not pink colored and control slides are stained as they should be...=20
Slightly orange color is probably produced by impuritys in the dye (acrid= in orange)... Some authors call for "basic fuchsine for schiff's" or "guaranteed acridin orange free pararosanilin" instead of basic fuchsin.
Basic fuchsin should be DISSOLVED COMPLETELY before adding the sodium metabisulfite and the HCl. A lot of recipes for Schiff's call for solutio= n of 1 gm of basic fuchsin in "HOT WATER" (80 - 100=B0C). Be sure to CLOSE = THE CONTAINER TIGHTLY after adding the sodium metabisulfite and the HCl and let stand for a few hours (some auteurs say "18 -24hrs") IN THE DARK. (Sorry: I'm not shouting, only emphasizing...).
The solution should have a strong "sulfur" smell.
I never had problems in preparing Schiff's. I use Basic fuchsin Merck "Certistain" and reagents UCB "PA" grade...
Several years a go our lab switched from the traditional KPTA to NHPTA. which enhances the fine structural details often lost with more aggressive stains. Prepare a 1% solution of phosphotungstic acid , then pH the stain with 1N ammonium hydroxide to 6.5 and / or 7.0. Regards, Skip
If you can't find a suitable resist for the KOH solution, I suggest dimpling the back (the Si) side most of the way through and then etching for a short time in KOH till the remaining thin region of Si is gone, but the surrounding thick support layer is mostly intact.
This should work very well in theory, though I've never done it myself.
For dimplers, if you have a choice, I recommend the D500i (or subsequent models) from the VCR Group, Inc.. It has exquisite thickness control, very good damping against vibration (invaluable with brittle samples), imbedded diamond abrasive wheels, and can be set up to produce a large flat bottomed dimple. (I've evaluated all makes before our purchase, and the VCR unit is by far the best - I have no interest or relationship with the manufacturer.)
If you have difficulty locating a dimpler, you can try ours, but we are short of man-power so you would have to travel and fiddle with it on your own. A better alternative might be to talk to the VCR applications lab and have them make a few samples for you as a way of demonstrating the instrument's capabilities. When I initially dealt with them, they were very accommodating. Unfortunately, I do not have the URL for the company, but an internet search should be fruitful with that.
An alternative technique might be a chemical jet polish, where the area of sample impacted by the jet of the etching solution thins preferentially. But that's a lot more tricky to control, and your remaining insoluble film must be able to withstand the pressure of the jet.
Whichever way you succeed, I would be interested what method(s) ended working for you.
Best of luck,
Valdemar rwafu-at-bsco.com or valdemar-at-fast.net
Valdemar Furdanowicz, Ph.D. Bethlehem Steel Co. Research G-165 Bethlehem, PA 18016-7699
Bethlehem -----Original Message----- } From: Edward Hirsch {edhirsch-at-att.net} To: Microscopy-at-sparc5.microscopy.com {Microscopy-at-sparc5.microscopy.com}
I have a Tracor Northern 8502 (5500/5700 hybrid) imaging system which is coupled with an ISI (now Topcon?) WB-6 SEM. I would like to transfer = the x-ray dot maps, x-ray spectra, and SEM images from the TN8502 to a=20 Win98 PC (TIFF, BMP, etc...). The 5700 side (imaging) runs on = Microware's=20 OS-9 and the 5500 (x-ray) side runs on a modified Fortran called = "flextran".
Please let me know if you have already solved this problem! Thanks in=20 advance for any suggestions.
Brian Reid Department of Chemistry University of Texas at Dallas 972-883-2709 breid-at-utdallas.edu
{!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} I have a Tracor Northern 8502 = (5500/5700 hybrid)=20 imaging system which is {BR} coupled with an ISI (now Topcon?) WB-6=20 SEM. I would like to transfer the {BR} x-ray dot maps, x-ray = spectra,=20 and SEM images from the TN8502 to a {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} Win98 PC (TIFF, BMP, etc...). = The 5700=20 side (imaging) runs on Microware's {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} OS-9 and the {/FONT} {FONT = color=3D#000000=20 size=3D2} 5500 (x-ray) side runs on a modified Fortran called=20 "flextran". {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} Please let me know if you have = already solved=20 this problem! Thanks in {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} advance for any = suggestions. {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} Brian Reid {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} Department of Chemistry {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} University of Texas at = Dallas {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} 972-883-2709 {/FONT} {/DIV} {DIV} {FONT color=3D#000000 size=3D2} {A=20 href=3D"mailto:breid-at-utdallas.edu"} breid-at-utdallas.edu {/A} {/FONT} {/DIV} {DIV} {FONT color=3D#000000 = size=3D2} {BR} {/FONT} {/DIV} {/BODY} {/HTML}
I'm no expert but we have had problems because of the following:
My experience with UA is that when it works it's great but it doesn't always work.
With PTA a bad batch of PTA stock could result in poor images - especially if it's old or not really a good quality reagent..
Do you adjust the pH of your PTA? It is normally fairly acid and should be adjusted to between 6 and 7 but you can experiment. We have always understood that it is best to use potassium hydroxide and not sodium hydroxide to adjust - personally I've seen little difference.
If you are using carbon coatings on your grids and clean preparations of virus maybe you need a little BSA to help spreading (not normally a problem with our samples).
If none of the above work it might be worth trying some of the newer more expensive stains like sodium silicotungstate or methylamine tungstate which can give nice results.
I suspect you may have tried most of the above, but I hope that something will help. Good Luck.
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD Tyne and Wear UK
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk
Disclaimer - all my own experiences and thoughts and it's mainlty black magic anyway,
---------- } From: Tom Phillips To: Microscopy
I have gotten some pretty good TEM's of a negative stained bacteriophage prep but, unlike some published photos of T4 and H. influenze phage, I don't get any fine details like striations in the tail. Essentially I get nice looking hexagons sitting on top of tails but no fine structure. I have tried staining with 1% PTA or 2% uranyl acetate or vandium. I have tried 2 basic protocols:
1. putting a drop of the prep on a carbon coated grid for 30 to 120 sec, wicking the drop off, rinsing with a drop of water, and then adding a drop of stain and almost immediately removing it. Skipping the rinse doesn't seem to help nor does lengthing the staining time.
2. putting a 5 ul drop of the prep on the grid and then adding a 5 ul drop of stain.
Are there any negative staining experts out there with hints for getting the best image?
TIA, tom
Thomas E. Phillips, Ph.D. Associate Professor of Biological Sciences Director, Molecular Cytology Core Facility
3 Tucker Hall Division of Biological Sciences University of Missouri Columbia, MO 65211 (573)-882-4712 (voice) (573)-882-0123 (fax)
by tartan.richmond.edu (8.8.8/8.8.8) with ESMTP id KAA16915 for {Microscopy-at-MSA.microscopy.com} ; Fri, 13 Nov 1998 10:15:09 -0500 (EST) X-Sender: gradice-at-facstaff.richmond.edu Message-Id: {l03130304b271fce4d6c6-at-[141.166.55.15]} Mime-Version: 1.0 Content-Type: text/plain; charset="us-ascii"
Thanks all for your tips on preparing Schiff's reagent. Here are most of the suggestions, except for a couple I inadvertently deleted. My original problem was that my protocol said the reagent should be clear, while mine remained colored.
I tried making up the Schiff Reagent for the first time a few weeks ago and had some problems getting it to turn out as the book said it should (I was using Hayat 1993, but the recipe is the same). My problem was that Hayat said that after shaking overnight, the solution should be clear tan colored; it instead went from grape juice to wine colored. My Prof. said just to go ahead and add carbon (I used animal charcoal) and filter. The resulting solution was a clear pale pink. He said that was okay, just add a pinch more metabisulfite and cap tightly. I did and the staining worked well. He says the metabisulfite is the important thing.
} From: hank p adams {hpadams-at-bcm.tmc.edu}
Gary, that is not an uncommon problem. It depends on your source of basic fucshin which is if my memory serves me right is a combination of pararosaline dyes. Some sources are better than others and I went thru quite a few. I had absolutely perfect results with some produced from Sigma under pararosaniline in their catalogue, though I don't remember which salt I used. The solution came out perfectly clear and I stored aliquots at minus 20C and it seemed quite stable.
} From: bozzola-at-siu.edu (John J. Bozzola)
Gary, It has been awhile since I worked with the Schiff reagent but I used to mix up large batches of it. Firstly, the formula that I have (Lillie (1954) is slightly different:
basic fuchsin (C.C.) 1 gm (best to use a dye certified by the Biological Stain Commission) distilled water 80 ml NaHSO3 is 2 gm or 1.9 gm Na2S2O5 N HCl 20 ml
I noticed that this formula uses more bisulfite and more HCl -- both of which generate more SO2 for decolorization of the fuchsin.
This should go in a tightly stoppered container (to keep in the SO2 which does the decolorization of the fuchsin) and shaken every 10 minutes for a period of 2 hr. Add 500 mg finely powdered fresh charcoal (we used coconut charcoal). Charcoal must be finely ground (like talc). You can do this twice to remove any residual color. Sometimes I even added charcoal, shake and store in a refrigerator overnight. Filter through several layers of paper filter media. When colorless store at 5 deg C. If white precipitate shows up, it can be refiltered. Pink color means time to discard it.
Since your stain is not dissolving, something may be wrong with the fuchsin. Lack of destaining may be caused by not enough bisulfite or loose fitting container. Most often, we did not even need to use the charcoal.
} From: "C. John Runions" {cjr14-at-cornell.edu}
Brian asks ...
} ... I would like to transfer the } x-ray dot maps, x-ray spectra, and SEM images ...
I don't believe your system would use any brand of file compression, so the bitmap should be intact. If you know the bitmap's size ... e.g., 512by512by8bit, then a program which can open a "raw" bitmap will work .. e.g., "Photoshop-} File-} Open as" ... indicate "raw" and enter the bitmap size and have it guess at the header size. Because some filetypes might also include info at the end of the file, you may have to guess at the header size yourself 'til the bitmap rows and columns align properly.
... 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/
John, 1:4 HF:Nitric will do what you want. It etches SiO2, but slowly. As long as you have more than a hundred Angstroms or so of SiO2, you should be able to remove the substrate with no trouble. It will help if you mechanically thin from the back to 100 um or so; the solution should get through the silicon in under 5 mins. Sample agitation / rotation will help keep the surface smooth, although this isn't really important for your application since it will smooth anyway when it reaches the SiO2.
Cheers,
Richard Beanland GMMT Ltd., Caswell, Towcester, Northants NN12 8EQ UK } } Hi, } } I was wanting to know if anyone has used any wax or etch resist } which will protect against warm KOH. I need to mask a Si/SiO2/Si } wafer (Silicon on Insulator) to preferentially remove a the thick Si } backside of my specimen and leave the surface Si/SiO2 bilayer intact } for TEM imaging. I was going to use warm KOH (50 C) to remove the Si } but I need to mask the surface and define a window on the backside Si } to etch through (the KOH will stop at the SiO2 layer). Any } suggestions? } } John } --------------------------------------- } John Michael Glasko } Materials Science and Engineering Dept. } 2142 Burlington Labs/Yarborough St } Raleigh, NC 27695-7916 } USA } tel: (919)-515-7217, fax: (919)-513-1699 } jmglasko-at-unity.ncsu.edu
There is a program from SAMx called TN2Win that we bought for this. It = will convert the spectra from TN format to ASCII x,y and EMSA format. I = forget the formats that the image files are stored in. There are some ramifications that you have to go through to do this. You have to have = a serial card on the TN5500 you have to boot the system to a remote = terminal which will be the PC. There are some cables that come witht he system = and both PC and TN software disks. The system was about $3500 if I = remember correctly. You can get info from their web site.
-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: Brian Reid To: Microscopy-at-Sparc5.Microscopy.Com
Dear Michelle,
It is possible that your protein is present in very little amount in the plant tissue. You may try the same fixation protocol for immunocytochemistry but embed in paraffin. Make thick sections, label it and look at it under a fluorescent scope. You may use a confocal scope too and optically slice the tissue to see if you get any labelling at all. If you use a confocal scope then use Cy5 as a fluorochrome, it gives very little background fluorescence in leaf tissue.
Good luck,
Soumitra Ghoshroy Ph.D. Department of Plant Sciences University of Arizona 303 Forbes Building Tucson, AZ 85721 Tel: 520-621-1230 Fax: 520-621-7186 e-mail: ghoshroy-at-ag.arizona.edu
On Wed, 11 Nov 1998, Michelle L. Peiffer wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Sorry that I didn't provide all the details on the original post, here they } are: } } We are attempting to label g-protein in Arabidopsis guard cells. Westerns } indicate the protein is water soluble, but also associates with membranes, } though it is not an integral membrane protein. Conventional fixation of } the cells, with aldehydes, osmium, embedded in Spurrs yeilds good } ultrastructure, but no labelling. For labelling we are fixing with 4% } paraformaldehyde, 0.5% glutaraldehyde in 0.1 M phosphate buffer; } dehydration in ethanol, embedding in LR White. The primary antibody is } polyclonal, affinity purified IgG, the secondary is goat anti-rabbit } conjugated to 10 nm colloidal gold. We are getting acceptable } ultrastructure and labelling in E. coli (expressing the protein) processed } this way, but not even a hint of labelling in plants. } } We do have the resources to do cryo-work, but the equipment is all brand } new, and we are still working out the details. Any suggestions on } improving this protocol will be greatly appreciated. Thanks } } } } #################################################### } Michelle Peiffer } Electron Microscope Facility for the Life Sciences } The Biotechnology Institute for Research and Education } 1 South Frear Lab } University Park, PA 16802 } 814-865-0212 email:mlk101-at-psu.edu } #################################################### } } }
Institution x Felix d'Herelle Reference Center for Bacterial Viruses, Department of x Microbiology, Faculty of Medicine, Laval University, Sainte-Foy, Canada. x
Title x A catalogue of T4-type bacteriophages. [Review] [115 refs] x
Source x Archives of Virology. 142(12):2329-45, 1997.
Sara E. Miller, Ph. D. P. O. Box 3020 Duke University Medical Center Durham, NC 27710 Ph: 919 684-3452 FAX: 919 684-8735
In a message dated 98-11-11 10:13:55 EST, mlk101-at-psu.edu writes:
{ { We are attempting to label g-protein in Arabidopsis guard cells. Westerns indicate the protein is water soluble, but also associates with membranes, though it is not an integral membrane protein. Conventional fixation of the cells, with aldehydes, osmium, embedded in Spurrs yeilds good ultrastructure, but no labelling. For labelling we are fixing with 4% paraformaldehyde, 0.5% glutaraldehyde in 0.1 M phosphate buffer; dehydration in ethanol, embedding in LR White. The primary antibody is polyclonal, affinity purified IgG, the secondary is goat anti-rabbit conjugated to 10 nm colloidal gold. We are getting acceptable ultrastructure and labelling in E. coli (expressing the protein) processed this way, but not even a hint of labelling in plants.
We do have the resources to do cryo-work, but the equipment is all brand new, and we are still working out the details. Any suggestions on improving this protocol will be greatly appreciated. Thanks } } Hi Michelle,
If the protein is water soluble, cryoultramicrotomy is probably going to be the solution to your problems. This doesn't fully explain why you can label the protein in E. coli that are expressing it. Perhaps the bacteria are producing so much of the protein that by the time they are processed and embedded there is still enough left in a conformation in the cells that it can be resolved by immuno. If the protein is in much lower concentrations in the guard cells the processing may cause its complete extraction or denaturation.
Until you can get comfortable with the cryo setup, perhaps you could try either quick freezing and freeze-drying or chemical fixation followed by low temperature embedding in a polar resin like Lowicryl K4M. The low temperatures combined with a polar resin might prevent some of the extraction and denaturation. Lowicryl is available from Energy Beam Sciences (1-800-992-9037).
If you freeze-dry the specimens you can do a quick vapor fixation with osmium tetroxide in a sealed bell jar or chamber. Under these circumstances the osmium seems to bind to the specimens in a different manner than in aqueous chemical fixation, and it does not go through the "secondary osmium black" formation like it does when it is dehydrated with ethanol. But it will have a fixative effect and help with the ultrastructure. If the specimens are fixed with osmium vapor you should probably avoid Lowicryl as the resin, since the black color will interfere with Lowicryl's polymerization by UV light.
We have shown in several publications that you can quick freeze and freeze-dry (or "molecular distillation" dry) erythrocytes, follow this by vapor fixation with aldehydes and osmium, embed in Spurr resin ( ! ) and still get excellent labeling of carbonic anhydrase, which is notoriously water solubile *and* fixative-labile.
One note of caution: If you vapor fix with osmium in a partial vacuum be sure the vacuum pump has a cold trap on it. If you don't have a cold trap be sure to change the pump oil after the run. The osmium will get into the oil and turn it black. This is a very effective, but somewhat expensive, osmium trap!
If you aren't set up for freeze-drying or Lowicryl embedding or don't want to go to the trouble, perhaps there is someone on this listserver that does such things on a regular basis and would be willing to help. But cryoultramicrotomy is really the way to proceed with this, imho.
Good luck, let us know how things turn out.
Best regards,
Bob **************************************** Robert (Bob) Chiovetti Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives Systems Integrators Analog & Digital Imaging *****************************************
I was given an assignment to map sulphur using EELS, does anyone have any good tips to map sulphur? Like microscope conditions, sample thickness, energy spread, etc. I've been trying to do it several times, but couldn't see any sulphur edge. Although, I detected sulphur using EDX.
I'm using FEG-CM20 with the Gatan PEELS.
3.81KV extraction voltage 3mm or 5mm PEELS apperature Gun lens 5 Spot size 6. energy dispersion 0.5 or 0.3 ev/channel.
I can't see any sulphur edge, but after the power law background substraction in EmiSpec there seems to be a variation of the sulphur I was mapping, but my advisor doesn't feel comfortable with the data because the entire spectrum seems to be varying ... so I'm not sure if it was from the thickness variation or the actual sulphur variation in the mapping. Are we supposed to see that actual sulphur edge? ... The sulphur content within the sample I'm looking at is not that much.
Please advise me ... any tips would be greatly appreciated ... I seem to be loosing hope in seeing this sulphur edge. Sometimes I even get delusional in seeing these edge ...
This international conference will focus on the latest developments in the study of the structural and electrical properties of semiconductors by the application of transmission and scanning electron microscopy, scanning probe microscopy and X-ray techniques.
The state-of-the-art in all important subject areas will be addressed, including the characterisation of bulk and thin film as-grown materials, the study of lattice defect and impurity behaviour and the investigation of advanced semiconductor processing procedures.
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The Proceedings of the conference will be published and the final call for papers has now been issued.
The abstract deadline is 4 DECEMBER 1998, and full abstract submission information can be found at the conference Web site http://www.iop.org/Confs. Enquiries may also be directed to Jacqueline Watts at the Institute of Physics, UK, Tel: +44-171-470-4800, E-mail: conferences-at-iop.org.
Dear Colleagues: Could anyone of you kindly tell me the titles and the publishers of good books on confocal microscopy? I need the ones for beginners or as more extensive reference book. Thanks in advance.
Yuhui Xu, MD,PhD Chief, EM Core Facility DFCI, Harvard Medical School
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Hello Tom,
In addition to the many recipes for stains you must be receiving, you = might also like to try a simple "low-dose" imaging protocol. I have found = when using negative stains that the image selection and focussing = procedures can damage fragile structures in the sample. Focussing on an = area close to a particle and then moving over to take the picture can help = prevent this. If you own a newer TEM, you might even have a low dose = capability built in. I have found this method very useful for small, = fragile samples, even when coated with thick films of aqueous uranyl = acetate or phosphotungstic acid.
Alternatively, the support film might be too thick to image the fine = detail. Is your grid carbon coated, as you post, or is it formvar/carbon = coated, which is more usual? Carbon alone offers more for high resolution = work.
Finally, have you checked out the vibrations affecting the microscope? = High resolution requires stable conditions.
Check out my web site for a brief summary of the options {http://www.hei.= org/htm/neg.htm} . =
I look forward to postings containing stain receipies. =
Paul Webster, Ph.D House Ear Institute 2100 West Third Street Los Angeles, CA 90057 phone:213 273 8026 fax: 213 413 6739 e-mail: pwebster-at-hei.org http://www.hei.org/htm/aemi.htm
by imo14.mx.aol.com (IMOv16.10) id LISBa17681; Fri, 13 Nov 1998 18:14:36 -0500 (EST) Message-ID: {6c2b6ab4.364cbd5c-at-aol.com}
Hi Tom,
I would only add to Paul Webster's and other's comments that an amorphous carbon film that has been glow-discharged in a partial vacuum with air for about 60 seconds is pretty hard to beat when it comes to a substrate for negative staining. We used this procedure for T4 on one of my (numerous) postdocs in the Dept. of Microbiology at the Biozentrum in Basel. We never found anything to beat it. The stain spreads beautifully on the surface.
Bob **************************************** Robert (Bob) Chiovetti Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives Systems Integrators Analog & Digital Imaging *****************************************
We have a Tracor ADEM SEM which runs on the OS9 software from Microware. We use a program called OS9MAX to convert TIFF images from the ADEM (8500) to PC readable format. The only glitch is you have to have a PC that has a 5.25" floppy. You save the image in OS9 TIFF format onto a floppy on the 8500, then you have to put that floppy in a PC running OS9MAX. The OS9 image is converted to PC TIFF readable and can be used in Photoshop or whatever. We have only saved images so far, I'm not sure we can convert spectra with the program.
If you are interested, I'll look up the name of the company we purchased OS9MAX from and send you more info. The company is in Germany but had a turn- around time of a few days between the time we made the purchase and received the software. E-mail me for more details.
THANK YOU SO MUCH for all the responses and advises (and possibly future response and advice as well). I think I should put them in a summary and put the advises into practice as soon as possible.
Roseann asked about the type of specimen I'm working with:
The sample is a biological sample ... looking for the sulphur content in the particle which gives us skin color.
Larry and Yasuo mentioned about confirming the sulphur with EDS (it's the same with EDX, right?) And whether the peak that I thought was sulphur could be coming from Mo or Pb. And about the beam damage.
Frankly, I don't know that Mo and Pb peaks should be there and I would have to look into that, but I doubt it is because if I do see Pb peak it would be another peak together with the sulphur. My 2 advisors think, it's definitely sulphur. "Sulphur concentration is not that much" and I think what you said about EELS sensitivity is absolutely correct .. and that's my problem, I think my sulphur edge is hidden in the tail of the plasmon. I have been trying to do 2nd difference spectra myself, but my EL/P computer just died (I was too harsh on it; so now, I would have to fixed the computer first before I could start working on 2nd difference again.)
And about the 2nd difference ... do you think MacFac by Bonnet and Trebbia would be an equally good solution as well?
I'm still really new in this field and haven't been doing much reading (that's my really bad part)... so I don't really know how thick my specimen is in term of mean free path but I'll find out soon ... but the physical thickness was 70nm and I just got a new one that is supposed to be 30nm thick.
As for beam damage, I try to work with the new areas to see the sulphur edge with EELS first, then if I can't see the edge (which is always the case) I would confirm it with EDS (which EDS always show the sulphur peak).
The assignment was given to me partially as a challenge to test my ability to use EELS. And to do EDS spectrum mapping would take a very long long time, so that's why my advisor wants me to do EELS spectrum mapping.
BTW, has anyone converted the EmiSpec file into EL/P file? Does anyone know where I could get the header file to convert Emispec file into EL/P file? I have the header file for Emispec file, but I don't know exactly where I could get the header file for EL/P. All the software in my lab is the first version (most primitive). :P
All-in-all THANK YOU so much ... and I would to put all the advises into good use. Below is Gerd's advice if anyone else is interested.
Thank you, Ad
+++++++++
} From Gerd:
First I would collect two spectra with good counting statistics in the sulphur and in the low loss region. Splice them and do the single scattering deconvolution using the LOG-ratio method in EL/P. This ensures that you get always the same background before the edge and you cancel out thickness effects.
Secondly, if you have regions without sulfur and without, you can use the spatial difference technique: author = {H. M\"ullejans and J. Bruley}, title = {Improvements in Detection Sensitivity by Spatial--Difference Electron Energy--Loss Spectroscopy at Interfaces in Ceramics}, journal = {Ultramicroscopy}, year = 1994, volume = 53, number = 4, pages = {351-360}, You need two spectra which are taken under the same conditions and at locations with the same thickness. Then you use the one without for an improved background subtraction.
I want to do freeze fracture and replication of bacteria and viruses. I am using the Balzers Freeze Etching System BAF 400. I managed to get to the state of floating off the replicas on a liquid surface. But this is accompanied by some problems and by using water they don't float off properly.
Should I use another solution for floating off the replica?
What about the influence of the way the specimen carriers have been cleaned?
There are different kinds of specimen carriers some do have a central boring and some do not. I have tried different kinds of combinations, but couldn't yet work out a difference. Are there any hints on this?
I ended up with quite a bit of ice on the cooled parts in the chamber which is floated with air. Is it highly recommended to use dry nitrogen instead?
Is there a special way to bake out the system after doing a freeze-fracture?
A final question for BAF specialists. Whenever the system switches to 'high vac' the valve located at the bottom of the chamber 'leaks' what means that it allows air to blow out (I think it's the Plate Valve PVA 160P). This doesn't effect the operation of the system, it just results in a nearly continuous working mode (+noise) for the compressor providing the high pressure for pneumatic system. Is this normal, or is there anything I can do against?
I am looking forward getting your replies!
Regards,
Bettina
*** Bettina Wolpensinger Electron Microscope Unit University of New South Wales Sydney, NSW 2052 AUSTRALIA phone: +612 9385 6390 fax: +621 9385 6400 b.wolpensinger-at-unsw.edu.au http://srv.emunit.unsw.edu.au ***
I believe that one of the other listers has already mentioned molybdenum peaks. This has been a chronic problem on our Hitachi H7000 because of the Mo in the movable objective apertures.
We have had problems with EDX, not EELS, but I would expect that the geometry may be worse in EELS because your detector might more directly "see" objective apertures and so increase the chance of picking up Mo. If your system will pick up higher energies of Mo it would certainly be worth doing a quick check.
Incidentally I received a lot of advice about our molybdenum problem some time ago and I can't remember if I thanked everyone - so thanks, just in case. Unfortunately all we ever managed to do was minimize the effect, unless we removed the aperture rod which produced its own problems.
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD Tyne and Wear UK
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk ---------- } From: Daraporn Arayasantiparb To: Microscopy
Dear Listers,
I was given an assignment to map sulphur using EELS, does anyone have any good tips to map sulphur? Like microscope conditions, sample thickness, energy spread, etc. I've been trying to do it several times, but couldn't see any sulphur edge. Although, I detected sulphur using EDX.
I'm using FEG-CM20 with the Gatan PEELS.
3.81KV extraction voltage 3mm or 5mm PEELS apperature Gun lens 5 Spot size 6. energy dispersion 0.5 or 0.3 ev/channel.
I can't see any sulphur edge, but after the power law background substraction in EmiSpec there seems to be a variation of the sulphur I was mapping, but my advisor doesn't feel comfortable with the data because the entire spectrum seems to be varying ... so I'm not sure if it was from the thickness variation or the actual sulphur variation in the mapping. Are we supposed to see that actual sulphur edge? ... The sulphur content within the sample I'm looking at is not that much.
Please advise me ... any tips would be greatly appreciated ... I seem to be loosing hope in seeing this sulphur edge. Sometimes I even get delusional in seeing these edge ...
by heinlein.acpub.duke.edu (8.8.5/Duke-4.6.0) with ESMTP id LAA03453; Mon, 16 Nov 1998 11:15:11 -0500 (EST) Received: (from saram-at-localhost) by godzilla2.acpub.duke.edu (8.8.5/Duke-4.6.0) id LAA19875; Mon, 16 Nov 1998 11:15:08 -0500 (EST)
James Pawley's Handbook of Biolobical Confocal Microscopy.
On Fri, 13 Nov 1998, Yuhui Xu wrote:
} Date: Fri, 13 Nov 1998 15:11:49 -0800 } From: Yuhui Xu {xuy-at-warren.med.harvard.edu} } To: Microscopy-at-sparc5.microscopy.com } Subject: Books on Confocal microscopy } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Dear Colleagues: } Could anyone of you kindly tell me the titles and the publishers } of good books on confocal microscopy? I need the ones for beginners } or as more extensive reference book. } Thanks in advance. } } Yuhui Xu, MD,PhD } Chief, EM Core Facility } DFCI, Harvard Medical School } }
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 want to do freeze fracture and replication of bacteria and viruses. I am } using } the Balzers Freeze Etching System BAF 400. I managed to get to the state of } floating off the replicas on a liquid surface. But this is accompanied by some } problems and by using water they don't float off properly. } } Should I use another solution for floating off the replica?
First you digest the sample, THEN you rinse with water. The "standard" is household bleach, but many samples will require something stronger. Have you read one of the basic books? } } What about the influence of the way the specimen carriers have been cleaned? } } Are there any hints on this?
Replicas usually are mounted on grids at the end of the cleaning process. Try moving the replica thru the sequence of cleaning solutions on a small glass ball. Make it by melting the tip of a disposable pipette. The replicas should float well. } } I ended up with quite a bit of ice on the cooled parts in the chamber which is } floated with air.
Not unusual; work faster.
Is it highly recommended to use dry nitrogen instead?
It helps!
} the valve located at the bottom of the chamber 'leaks' what means that it } allows air to blow out (I think it's the Plate Valve PVA 160P). This } doesn't effect the operation of the system,
Oh yes it does...
} Is this normal, or is there anything I can do against? } Get your system serviced! A Balzers specialist isn't necessary; anyone who understands vacuum systems will do. Ask your EM service person. }
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
Readers. Re: the microscopits salary survey that we have been conducting, I regret to advise that only some 500 folks supplied their salary data. So, as we breakdown to education and experience (even in 3 year increments), there simply is not enough data in each category to provide any reasonable results. It seems that we would have needed twice the participation to come up with useful final data. Many thanks to all who contributed in our effort. Don Grimes, Microscopy Today
A while back there was a thread on the safe range of solvents to use when cleaning a polymer thin window. I volunteered to test more solvents if requested.
The only one requested was hexane. We succesfully used hexane to clean a window, and went further to soak it for 20 minutes (not recommended!) without causing degradation in the leak test. However after soaking there was a slight residue visible under the microscope. This was likely caused by contamination.
best regards mark
Mark W. Lund, PhD VP Engineering } } 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
"This is a YOUNG business...How can I tell you what YOUR job is when I don't know what MINE is?" --Pogo
I don't know what responses were made to Daraporn off the listserver, but there seems to be some confusion in what has been posted on the server.
Daraporn has seen a small peak at the S energy in his EDX spectra, but could not find S in the EELS spectrum. This could be for one of two reasons: 1) the S is not S at all, but Mo or Pb. In this case, with luck, he might expect to see the K-lines of Mo or the L-lines of Pb at higher energy, provided, of course, that there is enough present (the low-energy lines at the S energy are much more intense than the higher energy ones) Mo could certainly be coming from apertures, as Malcolm Haswell suggests. Pb could, conceivably, be coming from the x-ray shielding in the microscope, but do other CM20's have that problem? There is no obvious reason why Daraporn's should if others don't, unless they have made some modification to the stage area. It would seem unlikely that Mo or Pb were actually in the sample. EELS would be used to confirm the analysis because EELS is not susceptible to "hole count" effects. To use the Mo aperture as an example: the Mo x-ray signal is generated by stray radiation (scattered electrons, x-rays) hitting the large area of the mostly thick aperture material. Hence just a few electrons or x-rays can produce lots of Mo x-rays. In contrast, the few electrons that penetrate the aperture would have lost lots of energy and been scattered through large angles, so would not reach the EELS detector at all. There is no equivalent of the mechanism that generated the spurious x-ray signal.
Unfortunately, there is another possible reason for not seeing the S, even if it is really there. The sample could be too thick. The whole relationship of beam current, sample thickness, statistics and spatial resolution is very complex, both in EDX and EELS analysis. However, one point is probably worth mentioning: Comparisons of the detection sensitivities for EDX and EELS compare the results from the same samples. Typically, in practice samples suitable for EELS are thinner than those often used for EDX analysis. Hence, although theory might say that (at least up to Ca or so) the detection limits for EELS are better than those for EDX, this is only true for thin samples. In thicker samples, the EELS backround rises and the peak becomes blurred out, resulting in poorer detection, while in EDX you just get more signal and hence better detectability.
The papers (from NIST/NBS) that illustrate the sensitivity of EELS also point out the need for extremely high incident beam currents, available only in a LaB6 microscope. I don't know enough about the CM20 FEG to translate from the settings that Daraporn used to actual beam characteristics. However, it is clear that the optimum beam settings for EELS are very different from those for EDX, and also the spatial resolution is significantly degraded. It could be that his microscope conditions are not close to optimum for EELS.
Hi, This is a bit off the microscopy subject - my lab has a Dahle paper trimmer 551 and someone has damaged the blade. This has been a great little cutter for many years. I'm looking for information on a replacement (blades or the whole thing). Most of the microscopy companies sell the Rotatrim cutters...any opinions and product suggestions are appreciated. thanks, Beth
************************************** Beth Richardson EM Lab Coordinator Botany Department University of Georgia Athens, GA 30602
You're right, I can save the file into text file first to get the header file, thanks.
Dr. Thomas, you gave me a very tough question because I really don't understand many of the terms that was used (it's really my fault, cause I haven't been reading much)... please don't brush me off yet.
I don't know what is the approx. S-content ... they just told me "it's low" .. but with my parallel probe analysis with 100sec collection time, meter plate reading of 2.5 sec ... the maximum sulfur peak with EDS was ~1000 counts. But when I switch to do the analysis in STEM mode, because the probe is small, the sulfur peak has to be collected with a longer time. While with EELS, I only need 5 sec or something like that.
I need the mapping mainly for the cosmetic purpose and because it's give us spatially resolved information, something like that. That's the term they used.
You mentioned about "point spread in the detection system" in DigiPEELS, what is that? No one mentioned about that. Is that in the newer version?
Could you explain briefly how I could optimize EDS acquisition? What are the things to look for? I think I tried to have the shortest data acquisition time and high beam current already. Do you think 2.5 sec meter plate reading is still low? I'm really not too familiar with this.
_____________
Mo peak ... to Dr. Garber and Dr. Malcolm
I don't know about Mo aperture, but I'll check ... but I think the microscope I'm using has Be-window for EDS, is that the movable Mo-aperture you are talking about. Or is it the aperture of the microscope?
Thank you so much for all the responses, Ad Daraporn
We are interested in studying HE (high explosive) materials by SEM. Typical nonsenstive HE are HMX (octahydro-1,3,5,7 tetranitro-1,3,5,7 tetrazocine), RDX, TATB, PETN etc. Do you have any experience with or know of any literature, operating procedure, guidelines? Any contact? Please respond or call as appropriate? Thank you. Tri Tran Energetic Materials Section, Lawrence Livermore National Lab (925) 422-0915 tran4-at-llnl.gov
I agree with Sara. We bought a new Rotatrim a year ago and it works great.
Patty Jansma Tel:520-621-6671 plj-at-manduca.neurobio.arizona.edu Arizona Research Labs Division of Neurobiology University of Arizona
On Mon, 16 Nov 1998, 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, } This is a bit off the microscopy subject - my lab has a Dahle paper trimmer } 551 and someone has damaged the blade. This has been a great little cutter } for many years. I'm looking for information on a replacement (blades or the } whole thing). Most of the microscopy companies sell the Rotatrim } cutters...any opinions and product suggestions are appreciated. } thanks, } 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 } ************************************** } } }
If anyone needs a large green premiere paper cutter I have one. It is too big to use for what I do and would swap it for microscope goodies or books on microscopes or microtechnique the bummer is that you have to pick it up in Arizona it is a monster!
Does anyone have the website or e-mail address where I can find JEOL's y2K compliancy statement regarding the JEOL 5400 - scanning electron microscope. I know that the software is not compliant and have ordered the upgrade. I am however concerned about date related functions in the embedded system.
We are looking for spherical beads (metal, glass, etc) with a well defined diameter (range : 1 - 500 micron) and a small standard deviation. These beads will be used for testing particle analysis software in SEM.
The equipment we use is a Philips XL 30 FEG SEM and Noran Voyager software.
Any suggestion where to get these beads will be welcomed.
Thank you very much in advance.
Please respond directly to :
vanderwulp-at-pml.tno.nl
Kees van der Wulp Prins Maurits Laboratory - TNO POBox 45 2280 AA, RIJSWIJK (ZH) Netherlands
-----Original Message----- } From: Tri Tran [mailto:tran4-at-popsicle.llnl.gov] Sent: Monday, November 16, 1998 6:38 PM To: microscopy-at-sparc5.microscopy.com
We are interested in studying HE (high explosive) materials by SEM. Typical nonsenstive HE are HMX (octahydro-1,3,5,7 tetranitro-1,3,5,7 tetrazocine), RDX, TATB, PETN etc. Do you have any experience with or know of any literature, operating procedure, guidelines? Any contact? Please respond or call as appropriate? Thank you. Tri Tran Energetic Materials Section, Lawrence Livermore National Lab (925) 422-0915 tran4-at-llnl.gov
For the interest of microscopists in KwaZulu-Natal and beyond:
One of our overseas guests to the MSSA conference in December, Dr Alice Warley, will be visiting the Centre for Electron Microscopy at the University of Natal in Pietermaritzburg during late November. During her visit with us she will be conducting an informal workshop on the preparation of standards for EDX in the TEM. She will also present a talk entitled "Elelements inside Cells - Qualitative and Quantitative X-ray Microanalysis in Biology'.
We may be able to accommodate a very limited number of extra persons for the workshop. All interested persons are welcome to attend the talk at 15h00 on Thursday, 26 November.
Tony Bruton Head, Centre for Electron Microscopy University of Natal, Pietermaritzburg Private Bag X01, Scottsville 3209, KwaZulu-Natal, South Africa. Tel +27 (0)331 260 5155, Home +27 (0)331 962676 Fax +27 (0)331 260 5776 email: bruton-at-emu.unp.ac.za
Along these lines, we've just received a copy of a short booklet which outlines a number of tests addressing the Y2K problem. Further infomration is available from Info Check, LLC in Manassas, VA. Price: $19.95. We have an aol address for them at CMontana4-at-aol.com. (MME has no commercial interest in this endeavor).
Your best bet for this is an art supply store, or the UG art departments. If UG has a printing office, try them. Most of these cutters are built for printers and artists.
Phil
} Hi, } This is a bit off the microscopy subject - my lab has a Dahle paper trimmer } 551 and someone has damaged the blade. This has been a great little cutter } for many years. I'm looking for information on a replacement (blades or the } whole thing). Most of the microscopy companies sell the Rotatrim } cutters...any opinions and product suggestions are appreciated. } thanks, } 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 } **************************************
}}}}}}}}}}}}}}}}}}}}}}}}}}{{{{{{{{{{{{{{{{{{{{{{{{{{ Philip Oshel PO Box 620068 Middleton, WI 53562 oshel-at-terracom.net
I would be grateful if you would bring the available faculty position below to the attention of potential candidates.
thanks,
Nigel Browning
Faculty Position University of Illinois at Chicago
The Department of Physics at the University of Illinois at Chicago (UIC) is planning to fill one tenure-track assistant professorship position to begin Fall 1999. The department has a strong commitment to both undergraduate and graduate education and is dedicated to excellence in research. At present, we have thirty faculty members active in many areas of physics and biophysics. Our search will give special consideration to candidates in the areas of condensed matter physics and biophysics.
Successful candidates will have a Ph.D. degree or equivalent, are expected to develop top quality research programs and should show high promise of obtaining external research funding. We encourage research programs which take advantage of the numerous facilities in the Chicago area. Applicants should submit a vita and a statement of research interests and future plans. They should also arrange to have three letters of recommendation sent to: Prof. Inder P. Batra, Head, Department of Physics, University of Illinois at Chicago (MC 273), 845 W. Taylor St., Rm. 2236, Chicago, IL 60607-7059. Applications will be considered as they are received, with a deadline of March 25, 1999. The University of Illinois at Chicago is an Affirmative Action/Equal Opportunity Employer.
The National Institute of Standards and Technology sells polystyrene be= ads designed for particle sizing (SRM 1692). Information is available at http://ts.nist.gov/srm.
Joe Neilly Abbott Laboratories Dept. of Microscopy and Microanalysis Abbott Park, IL 60064-3537 =
Kees, Try the following. They may have what you need. We've purchased various microspheres from them with successful outcomes.
Bangs Labs, Inc. 9025 Technology Drive Fishers, Indiana 46038-2886 USA E-mail: info-at-bangslabs.com http://www.bangslabs.com
Good luck. Winston ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~ Winston W Wiggins, Supervisor 11/17/98 11:53:38 AM CRC-Electron Microscopy Lab. Ofc:704/355-1267 Carolinas Medical Center Fax:704/355-7648 P.O. Box 32861 Lab:704/355-7220 Charlotte,NC 28232-2861 USA Eml:wwiggins-at-carolinas.org ~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~
Does anyone have a procedure for distinguishing between hard and soft c= lay? Any information would be useful. Please don't hesitate to contact me o= ff line if you think this information would be of limited interest to the group=
Hair today, gone tomorrow I was recently asked by someone who develops hair color products, ( I didn't think I was greying that much) if I might be able to recommend a "system" that could be used to evaluate the efficacy of various hair colorants. Because this person would like to complete the examination without removing any hair, i.e. in-situ and in real time the "system" would have to be portable and consist of relatively high magnification optics coupled to a video camera with the signal fed to a color monitor. The desired magnification of such a system is 400 - 500 times. I am not familiar enough with the video camera market to identify/recommend a camera for such an application but am hopeful that someone out there can help.
Thanks, Paul Gerroir Xerox Research Centre of Canada
Is there a used SEM that requires relocation? We have few capital funds so we are looking for a second hand instrument that has out lasted its usefulness and needs a new home. If you have such a microscope within a reasonable distance to me and wish to negotiate terms and conditions of its removal, please contact me by e-mail below.
Best regards,
Gary
................................................................ Dr. Gary Faulkner, Ph.D. Director, Faculty of Medicine EM Facility Department of Microbiology & Immunology Sir Charles Tupper Building Dalhousie University Halifax, N.S. B3H 4H7 Tel: 902.494.2346 Fax: 902.494.5125 E-mail: Gary.Faulkner-at-Dal.Ca
How to improve the contrast in plant tissue included in Spurr resin? The double contrast with Uranyl acetate in ethanol solution and Lead citrate is not good for me. There is some special procedure for to solve this problem? With the ultra-low viscosity resin, the problem is worst (lower contrast than the Spurr resin). Thank's in advance.
M.Sc. Rinaldo Pires dos Santos Dept. of Botany - UFRGS Porto Alegre - RS - Brazil
with ESMTP id {01J4A9WJ7X9I8Y5EXU-at-denver.du.edu} for Microscopy-at-MSA.Microscopy.Com; Tue, 17 Nov 1998 15:30:50 MST Received: from localhost by du.edu (PMDF V5.1-10 #28062) with SMTP id {0F2L00N017W3YO-at-du.edu} for Microscopy-at-MSA.Microscopy.Com; Tue, 17 Nov 1998 15:31:15 -0700 (MST)
Hi,
We are having a problem retaining synaptic vesicles in tissue fixed with picric acid and 0.25% glut, embedded in LR Gold, and polymerized at -20 with UV. We will loose the antigen if we raise the percentage of glut. We cannot use osmium because we loose the antigen and then also cannot use UV for polymerization. Is there anyone who has been able to preserve vesicles with the above approach? Does anyone have any novel ideas? We will try tannic acid. Has anyone used this for vesicle preservation? I would so much appreciate any comments on what we are trying to do. Bye, Hildy {hcrowley-at-du.edu}
I really don't know how they made these slides back then. Check them out. It may be worth your while to look at these rare items..any info on how they were made would be appreciated. Thanks Dan Fleming
To view: http://cgi.ebay.com/aw-cgi/eBayISAPI.dll?ViewItem&item=42528374
This is not a forsale post but a request for information.
Frank - Please try the Home Page of the Clay Minerals Society: {http://shadow.agry.purdue.edu/clay/claymin/claymins.html} where you will find directions for subscribing to their list server. I'm sure someone there will be able to help you.
Being a clay mineralogist myself, I'll give it a try. Hard & Soft are commercial (common sense) terms meaning just that. Hard clays (such as "hard kaolin", also see "flint clay") require grinding and blending, wheras "soft kaolin" may require less preparation. Plastic clays are soft, wet and like modeling clay. Then there's "ball clay" which can be rolled up into a ball! Hard and Soft have no clear mineralogic or chemical meaning. See the AGI Glossary of Geology (1997?) for more info. Dave Pevear, Houston
FRANK KARL wrote: } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } Does anyone have a procedure for distinguishing between hard and soft clay? } Any information would be useful. Please don't hesitate to contact me off line } if you think this information would be of limited interest to the group
for Microscopy-at-Sparc5.Microscopy.Com id AA24217; Wed, 18 Nov 98 16:00:20 +0100 Message-Id: {9811181500.AA24217-at-iris1.el.ub.es}
Dear colleagues,
We are working on the TEM characterisation of CuInS2 (usually called CIS) on glass substrates. We are preparing TEM samples out of these materials (both plan view and cross-section) and we are encountering problems with the preparation procedure, which are strong amorphisation of the CIS layer in plan-view and strong damage (not always amorphisation) in cross-section.
The preparation procedure we use is the standard preparation method we use for Si-based materials: For cross-section, we cut stripes out of the samples, glue them together. Next flat grinding, dimpling and finally ion milling are used. For plan-views, a piece of sample is ultrasonic cut and the preparation continues with the flat grinding, ... and ion milling only from the backside.
We believe that the problems we have are strongly related to the glass substrate and to the charging of the glass during ion milling, resulting in an overheating of the sample. This leads to extremely poor cross-sections and to even worst plan-view samples (however we still have some good results from few of these samples). It might be also that the CIS layer is strongly beam sensitive (any experience with it?).
Any help in trying to circumvent these problems will be extremely helpful and experience in preparing samples of the type glass substrate-thin layer would also be strongly appreciated.
By the way our ion milling machines can work at liquid nitrogen temperature in order to minimise sample heating, if this helps.
Thank you in advance for your answers.
Albert Romano-Rodriguez Dept. of Electronics Faculty of Physics University of Barcelona c/ Marti i Franques, 1 E-08028 BARCELONA Spain tel: +34-93-402 11 47 FAX: +34-93-402 11 48 e-mail: romano-at-el.ub.es
If not, I suspect that what is for sale is not a series of slides but a series of plates (hand painted pages to insert into a book) made by painting what was observed on a slide.
Gerard Turner, at Oxford, manages the collection of the Royal Microscopical Society. Either he or Dr. Savile Bradbury (Oxford, retired), would probably be the best sources for information on this sort of thing. Here in this country, Cecil Fox, who had a lot of contact with the Billings Collection at the Armed Forces Institute of Pathology, might know. Also, Don O'Leary manages the collection for the New York Microscopical Society. Try him at 201-797-8849.
You can probably contact either Gerard or Savile by writing/emailing Oxford University in the UK. I only have old contact information for Cecil Fox but would be happy to provide that information off-line if you are interested in talking to him.
Friends, I am interested in contacting some one who does micro CT imaging. If you are one or know of one I would appreciate hearing from you.
Greg Erdos Gregory W. Erdos, Ph.D. Ph. 352-392-1295 Assistant Director, Biotechnology Program PO Box 110580 Fax: 352-846-0251 University of Florida Gainesville, FL 32611
Responding to the message of {3651F0D1.6C4F-at-botanica.ufrgs.br} from Rinaldo Pires dos Santos {rinaldop-at-botanica.ufrgs.br} : } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Dear colleagues } } How to improve the contrast in plant tissue included in Spurr resin? The } double contrast with Uranyl acetate in ethanol solution and Lead citrate } is not good for me. There is some special procedure for to solve this } problem? With the ultra-low viscosity resin, the problem is worst (lower } contrast than the Spurr resin). } Thank's in advance. } } M.Sc. Rinaldo Pires dos Santos } Dept. of Botany - UFRGS } Porto Alegre - RS - Brazil
What kind of plant tissues?
1. Try a 50:50 mixture of Spurrs/Embed812 resins. Mix up resins seperately without their accelerators (DMAE & DMP-30). You can store them in the freezer. Then mix 50:50 just before use, mix in required amounts of the two accelerators, infiltrate and cure. You should get the better sectioning and staining properties of Embed812 plus the low viscosity of Spurrs.
2. Switch to Embed 812 100% and extend infiltration times.
3. What lead stain are you using? In my experience, Reynold's lead citrate doesn't seem to work too well with Spurrs. I use Sato's triple lead stain, preceeded by 3% UA, and get very good staining on plant and bacterial samples embedded in Embed812.
Good luck,
Gib
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
Our group currently has a JEOL FX 2000 TEM available for sale to anyone interested. The system includes a cold and hot stage, PC driven X-ray system,and STEM attachment. Please respond if interested.
M.W. Rigler, Ph.D. MAS, Inc. Suwanee GA 770-866-3218
Return-Path: {Microscopy-request-at-sparc5.microscopy.com} Received: from relay29.mx.aol.com (relay29.mail.aol.com [172.31.109.29]) by air09.mail.aol.com (v51.16) with SMTP; Thu, 12 Nov 1998 08:47:16 -0500 Received: from Sparc5.Microscopy.Com (sparc5.microscopy.com [206.69.208.10]) by relay29.mx.aol.com (8.8.8/8.8.5/AOL-4.0.0) with SMTP id IAA00928; Thu, 12 Nov 1998 08:47:05 -0500 (EST) Received: (from daemon-at-localhost) by Sparc5.Microscopy.Com (8.6.11/8.6.11) id HAA07680 for dist-Microscopy; Thu, 12 Nov 1998 07:22:03 -0600 Received: from no_more_spam.com (Sparc5 [206.69.208.10]) by Sparc5.Microscopy.Com (8.6.11/8.6.11) with SMTP id HAA07677 for "MicroscopyFilteredEmail-at-msa.microscopy.com"; Thu, 12 Nov 1998 07:21:32 -0600 } From: "Mriglermas-at-aol.com"-at-sparc5.microscopy.com Received: from imo17.mx.aol.com ([198.81.17.7]) by Sparc5.Microscopy.Com (8.6.11/8.6.11) with ESMTP id HAA07670 for {Microscopy-at-sparc5.microscopy.com} ; Thu, 12 Nov 1998 07:21:13 -0600 Received: from Mriglermas-at-aol.com by imo17.mx.aol.com (IMOv16.10) id NYLNa04133 for {Microscopy-at-sparc5.microscopy.com} ; Thu, 12 Nov 1998 08:31:14 -0500 (EST) Message-ID: {d40385e2.364ae322-at-aol.com}
Our group currently has a JEOL FX 2000 TEM available for sale to anyone interested. The system includes a cold and hot stage, PC driven X-ray system,and STEM attachment. Please respond if interested.
M.W. Rigler, Ph.D. MAS, Inc. Suwanee GA 770-866-3218
To: Dan Fleming Re: Antique Microslides Dan, The expert on this subject is Dr. James B. McCormick. He has been assisted by M. Lamar Jones at many workshops during NSH conventions. You can get more info from National Society for Histotechnology at 301-262-6221.
Bob Santoianni Emory University Hospital Atlanta, Georgia robert_santoianni-at-emory.org
Kees - I've used 9.870 um (+/- 0.057 um) polystyrene spheres from Duke Scientific Corp with good success. I believe the company offers a large range in sizes as well. Their web site is www.dukescientific.com. Hope this is helpful.
Dave Joswiak University of Washington Dept. of Astrononmy Seattle, WA 98195 joswiak-at-astro.washington.edu
On Tue, 17 Nov 1998, Wulp, Cees van der wrote:
} ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } -----------------------------------------------------------------------. } } } Dear SEMmers, } } We are looking for spherical beads (metal, glass, etc) with a well defined } diameter (range : 1 - 500 micron) and a small standard deviation. } These beads will be used for testing particle analysis software in SEM. } } The equipment we use is a Philips XL 30 FEG SEM and Noran Voyager software. } } Any suggestion where to get these beads will be welcomed. } } Thank you very much in advance. } } Please respond directly to : } } vanderwulp-at-pml.tno.nl } } Kees van der Wulp } Prins Maurits Laboratory - TNO } POBox 45 } 2280 AA, RIJSWIJK (ZH) } Netherlands } }
I would suggest using a tripod polisher for getting the specimen to as thin as possible. This basically involves polishing the specimen to a thin wedge (between 0.5 - 3 degrees) using a set of polishing mats with varying grain size (30 um to 500 nm). I have polished silicon (mechanically) with this method to about 40 nm! But for most cases it will get you down to below 1um. The method uses a liquid to aid polishing lubrication, but you can use either oil based or water. I have a colleague who has used this to get his Barium based glass spcimens down to a couple of microns before finishing the polishing with a quick ion mill ( { 1 hr) and has report very good results. I have also used this specimen to map the Fe content in the crystallites in the glassy matrix.
The tripod polisher is produced by South Bay Technologies (sorry I do not have an address or phone) but you could try a web search.
I hope this helps. Yours, Jonathan Barnard Microstructural Physics, H.H.Wills Physics Lab. University of Bristol, U.K
Dear Paul, The last time I was at the MSC meeting, Chris Cathcart of Marivac had exactly the thing you are looking for, I believe. It was a little, portable video camera and lens system with built-in illumination. He held it up to his tie and we could see a brilliant image of the threads in the weave, in full colour. Contact him at Marivac, in Canada 1-800-565-5821. Good luck. You wrote:
} Hair today, gone tomorrow } I was recently asked by someone who develops hair color products, ( I didn't } think I was greying that much) if I might be able to recommend a "system" } that could be used to evaluate the efficacy of various hair colorants. } Because this person would like to complete the examination without removing } any hair, i.e. in-situ and in real time the "system" would have to be } portable and consist of relatively high magnification optics coupled to a } video camera with the signal fed to a color monitor. The desired } magnification of such a system is 400 - 500 times. I am not familiar enough } with the video camera market to identify/recommend a camera for such an } application but am hopeful that someone out there can help. } } Thanks, } Paul Gerroir } Xerox Research Centre of Canada } 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
The person that would probably be the most helpful to you is Scott Walck at PPG, who is pretty well the King of cleaved glass TEM samples (Walck-at-ppg.com). He has developed a variation of the small-angle cleavage technique for preparing outstanding cross-sectional samples of glass or thin films on glass. This is a non-charging, heat-free preparation technique. A good reference is:
Scott D. Walck and John P. McCaffrey, "The small-angle cleavage technique: an update", Mat. Res. Soc. Symp. Proc. Vol. 480, Materials Research Society (1997) pp149-171.
I have an extra copy and will mail it, along with an instructional video of the small-angle cleavage technique that Scott and I made .
Cheers John
John P. McCaffrey Institute for Microstructural Sciences National Research Council of Canada M-50 Montreal Rd. Ottawa, Ontario K1A 0R6 Canada
---------- } From: Albert Romano-Rodriguez To: Microscopy-at-sparc5.microscopy.com -----------------------------------------------------------------------.
Dear colleagues,
We are working on the TEM characterisation of CuInS2 (usually called CIS) on glass substrates. We are preparing TEM samples out of these materials (both plan view and cross-section) and we are encountering problems with the preparation procedure, which are strong amorphisation of the CIS layer in plan-view and strong damage (not always amorphisation) in cross-section.
The preparation procedure we use is the standard preparation method we use for Si-based materials: For cross-section, we cut stripes out of the samples, glue them together. Next flat grinding, dimpling and finally ion milling are used. For plan-views, a piece of sample is ultrasonic cut and the preparation continues with the flat grinding, ... and ion milling only from the backside.
We believe that the problems we have are strongly related to the glass substrate and to the charging of the glass during ion milling, resulting in an overheating of the sample. This leads to extremely poor cross-sections and to even worst plan-view samples (however we still have some good results from few of these samples). It might be also that the CIS layer is strongly beam sensitive (any experience with it?).
Any help in trying to circumvent these problems will be extremely helpful and experience in preparing samples of the type glass substrate-thin layer would also be strongly appreciated.
By the way our ion milling machines can work at liquid nitrogen temperature in order to minimise sample heating, if this helps.
Thank you in advance for your answers.
Albert Romano-Rodriguez Dept. of Electronics Faculty of Physics University of Barcelona c/ Marti i Franques, 1 E-08028 BARCELONA Spain tel: +34-93-402 11 47 FAX: +34-93-402 11 48 e-mail: romano-at-el.ub.es
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Reply to: RE: Synaptic Vesicle Preservation??????? Dear Hildy,
Are you sure your problem is with retaining the synaptic vesicles? Could = it be more a problem of contrast? As I know it is possible to visualize = synaptic vesicles in brain fixed with 4% formaldehyde alone, I will assume = your problem is contrast. Here are a few suggestions.
1. Check to see if your antigen is present and the antibody works. Try = the labeling by light microscopy first (cryostat sections will work well = for this). If you can't get a signal by light microscopy go back to = understanding more about your antigen and the antibody. =
2 Check LR White labeling by light microscopy. If you can get a signal = in cryostat sections then use the same fixation protocol to embed the = tissue in LR WHite. A good protocol is to fix the tissues in 4% = formaldehyde alone. If the LM labeling only works after methanol fixation,= try that for the EM too. Mount the resin-embedded sections on glass and = label them for light microscopy. Ideally, you should be able to compare = both the antibody labeling protocol you use for routine LM labeling with = as much of the EM labeling protocol you can use. Silver enhancement = protocols for colloidal gold are useful here. If you can't see a signal, = it may be that the LR White is affecting the antigen-antibody interaction. = There is no way you will be able to perform EM until the LM conditions = have been worked out.
3. Improving contrast in LR White. If you can get the antibodies to label = LR White sections lable the sections for EM. You should see a signal. If = you don't see a signal go back to step 2. If you do see a signal, the = antigen is there. If you don't see the structure that should label, then = your problem is with section contrast. There are many ways to manipulate = this. = a) cut thicker sections. That sometimes helps. = b) examine the section more. This will burn out resin from the section = and may help. c) extract the tissue prior to embedding. Although detergents, low = osmolarity fixatives and modified buffers can have the effect you need (= less cytoplasm) be careful that your antigen is not extracted too. = Knowing as much about the antigen as possible is useful here. d) treat the tissue with osmium tetroxide or uranyl acetate prior to = embedding. Who says osmium tetroxide will cause you to lose your antigen = or stop you using UV polymerization? A good way to incorporate = contrasting agents is to freeze substitute a frozen sample on dry ice in = medium containing up to 1% OsO4 or uranyl acetate. At the lowered temperat= ure there is no production of the black reaction product typical of osmium = treatment. Make sure that OsO4 is washed away while the tissue is still = on dry ice. Want a protocol? try this: {http://www.hei.org/htm/pmfs.htm} . = Be brave.
4. Use cryosections for immunolabeling. If after all your attempts to get = a positive signal you get nothing, it may be that the resin is blocking = the antibody-antigen interaction. This sometimes happens. It may be time = for you to check out the modern world of cryosectioning. Technology and = specimen preparation protocols have improved enormously over the last few = years and make cryosectioning an almost routine technique. Check it out: {= http://www.hei.org/htm/cryo.htm}
Please feel free to contact me off-line if you need clarification of these = points (I did leave out many details). I am still sitting in an empty lab = (no TEM, no ultramicrotome etc!) so have more time to spare than usual.
Regards,
Paul Webster, Ph.D House Ear Institute 2100 West Third Street Los Angeles, CA 90057 phone:213 273 8026 fax: 213 413 6739 e-mail: pwebster-at-hei.org http://www.hei.org/htm/apw.htm
HILDEGARD CROWLEY wrote:
} Hi, } } We are having a problem retaining synaptic vesicles in tissue fixed with } picric acid and 0.25% glut, embedded in LR Gold, and polymerized at -20 } with UV. We will loose the antigen if we raise the percentage of glut. = We } cannot use osmium because we loose the antigen and then also cannot use = UV } for } polymerization. } Is there anyone who has been able to preserve vesicles with the above } approach? } Does anyone have any novel ideas? We will try tannic acid. Has anyone } used this for vesicle preservation? } I would so much appreciate any comments on what we are trying to do. } Bye, } Hildy } {hcrowley-at-du.edu} } } } } } RFC822 header } ----------------------------------- } } Received: from Sparc5.Microscopy.Com [206.69.208.10] by mailhouse.hei.= org } (SMTPD32-4.07) id AEDC1EBF02A2; Tue, 17 Nov 1998 21:45:00 PST } Received: (from daemon-at-localhost) by Sparc5.Microscopy.Com (8.6.11/8.6.= 11) = } id QAA11965 for dist-Microscopy; Tue, 17 Nov 1998 16:39:58 -0600 } Received: from no_more_spam.com (Sparc5 [206.69.208.10]) by = } Sparc5.Microscopy.Com (8.6.11/8.6.11) with SMTP id QAA11960 for = } "MicroscopyFilteredEmail-at-msa.microscopy.com"; Tue, 17 Nov 1998 16:39:26 -= 0600 } Received: from atlas.cair.du.edu (atlas.cair.du.edu [130.253.2.202]) by = } Sparc5.Microscopy.Com (8.6.11/8.6.11) with ESMTP id QAA11947 for = } {Microscopy-at-MSA.Microscopy.Com} ; Tue, 17 Nov 1998 16:39:08 -0600 } Received: from odin.cair.du.edu by denver.du.edu (PMDF V5.1-12 #28064) } with ESMTP id {01J4A9WJ7X9I8Y5EXU-at-denver.du.edu} for } Microscopy-at-MSA.Microscopy.Com; Tue, 17 Nov 1998 15:30:50 MST } Received: from localhost by du.edu (PMDF V5.1-10 #28062) } with SMTP id {0F2L00N017W3YO-at-du.edu} for Microscopy-at-MSA.Microscopy.Com; = Tue, } 17 Nov 1998 15:31:15 -0700 (MST) } Date: Tue, 17 Nov 1998 15:31:15 -0700 (MST) } From: HILDEGARD CROWLEY {hcrowley-at-du.edu} } Subject: Synaptic Vesicle Preservation??????? } X-Sender: hcrowley-at-odin.cair.du.edu } To: postmessage {Microscopy-at-Sparc5.Microscopy.Com} } Message-id: {Pine.OSF.3.95.981117151619.28840A-100000-at-odin.cair.du.edu} } MIME-version: 1.0 } Content-type: TEXT/PLAIN; charset=3DUS-ASCII } Content-transfer-encoding: 7BIT } Errors-to: Microscopy-request-at-sparc5.microscopy.com } X-UIDL: 907881491 } Status: U } =
Paul Webster, Ph.D House Ear Institute 2100 West Third Street Los Angeles, CA 90057 phone:213 273 8026 fax: 213 413 6739 e-mail: pwebster-at-hei.org http://www.hei.org/htm/apw.htm
by post2.fast.net (8.8.8/8.8.5) with SMTP id NAA04968; Wed, 18 Nov 1998 13:45:05 -0500 (EST) Message-ID: {000d01be1323$3091d7c0$02f993cd-at-rfurdanowiczw.hrl.bsco.com}
Materials Characterization Facility University of Minnesota
The Center for Interfacial Engineering at the University of Minnesota is = seeking an instrument specialist as a staff member in its materials characterizat= ion =
facility. The facility houses 5 EM's, 6 XRD/SAXS instruments, several SPM= 's, =
micro-indentors, and other surface and thin-film analytical instruments. = See our website for details (http://resolution.umn.edu). The person will work mai= nly in =
the EM laboratories. The principle responsibilities of the position incl= ude =
training researchers to operate transmission and scanning electron micros= copes, =
maintaining and operating the TEM=D5s (Philips CM30 and JEOL1210), and as= sisting =
users in TEM specimen preparation and data interpretation. The position r= equires a Ph.D. in biosciences, materials science, physics or related discipline.= Very =
strong hands-on experience in various TEM techniques and their applicatio= n to =
materials characterization is required. Applicants should also have the =
experience and flexibility to work with other techniques. Experience in s= pecimen preparation and working in a multi-user facility is particularly desirabl= e. This is an annually renewable professional appointment; 12 month, 100% time re= gular =
appointment with excellent university benefits. Position and salary will= be =
commensurate with education and experience.
Please send resume, three letters of recommendation and salary requiremen= ts to =
Elizabeth Guldan, Search Committee, Center for Interfacial Engineering, =
University of Minnesota, 187 Shepherd Labs, 100 Union St. SE, Minneapolis= , MN =
55455. Screening will begin on January 31, 1999 and end when a suitable =
applicant is identified.
The University of Minnesota is an Equal Opportunity Educator and Employer= =2E
__________________ Stuart McKernan stuartm-at-tc.um= n.edu Microscopy Specialist Office:(612) 626= -7594 CIE Characterization Facility, University of Minnesota Desk: (612) 624= -6009 100 Union Street S. E., Minneapolis, MN 55455 Lab: (612) 624= -6590
Albert: I am not familiar with the mechanical characteristics of CIS, but I wonder if you would do better trying tripod polishing and maybe avoid having to do ion milling ??.
Jordi Marti
Dear colleagues,
We are working on the TEM characterisation of CuInS2 (usually called CIS) on glass substrates. We are preparing TEM samples out of these materials (both plan view and cross-section) and we are encountering problems with the preparation procedure, which are strong amorphisation of the CIS layer in plan-view and strong damage (not always amorphisation) in cross-section.
The preparation procedure we use is the standard preparation method we use for Si-based materials: For cross-section, we cut stripes out of the samples, glue them together. Next flat grinding, dimpling and finally ion milling are used. For plan-views, a piece of sample is ultrasonic cut and the preparation continues with the flat grinding, ... and ion milling only from the backside.
We believe that the problems we have are strongly related to the glass substrate and to the charging of the glass during ion milling, resulting in an overheating of the sample. This leads to extremely poor cross-sections and to even worst plan-view samples (however we still have some good results from few of these samples). It might be also that the CIS layer is strongly beam sensitive (any experience with it?).
Any help in trying to circumvent these problems will be extremely helpful and experience in preparing samples of the type glass substrate-thin layer would also be strongly appreciated.
By the way our ion milling machines can work at liquid nitrogen temperature in order to minimise sample heating, if this helps.
Thank you in advance for your answers.
Albert Romano-Rodriguez Dept. of Electronics Faculty of Physics University of Barcelona c/ Marti i Franques, 1 E-08028 BARCELONA Spain tel: +34-93-402 11 47 FAX: +34-93-402 11 48 e-mail: romano-at-el.ub.es
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Hi,
I wonder if I could solicit advice off-line about the benefits and = drawbacks of currently available critical point dryers and high resolution = sputter coaters? All replies will be treated as opinions and will remain = confidential. =
Regards,
Paul Webster, Ph.D House Ear Institute 2100 West Third Street Los Angeles, CA 90057 phone:213 273 8026 fax: 213 413 6739 e-mail: pwebster-at-hei.org http://www.hei.org/htm/apw.htm
Thanks to all responses. I think I have quite a number of things I would need to do and adjust and look into in my experiment before I could get back to all of you. All responses and tips and advises were very helpful. I'll put all of them into practice and pray for a better results this time.
Does anyone care for me to put all responses into a summary?
Special thanks to
Dr. Larry Thomas Dr. Malcolm Haswell Dr. Tony Garratt-Reed Dr. Adam Papworth Dr. Michal Jarnik Dr. Richard Leapman Dr. Charles Garber Dr. Yasuo Ito Dr. Roseann Csencsits and Dr. Gerd Duscher
} Dear Hildy } I have had good results with neural preservation-- fixing with 4% } formaldehyde (EM grade, no methanol) +.2% glutaraldehyde in PBS. } Dehydration was partial and in a methanol series with progressive } lowering of temperature, } infiltrated in LR white at -20C, and polymerizing at 55C. The series of } methanol was 60% for 15' at 4C, 70% methanol for 60' at -20C, and 80% } methanol for 60' at -20C, then 3 changes of LR white for 60' at -20C. Hope } this will help. This method has given good antigenicity with a wide } variety of tissues. If you need further info, please feel free to contact } me. } Marge } } Margaret Springett } e-mail hukee.margaret-at-mayo.edu } IEM Specialist at Mayo Foundation } 1426 Guggenheim } Rochester, Mn. 55905 }
Margaret Springett e-mail hukee.margaret-at-mayo.edu IEM Specialist at Mayo Foundation 1426 Guggenheim Rochester, Mn. 55905
A major development in ion milling is just now being released and may be applicable to your problem. Amorphous damage (as well as implanted Ga fr= om FIB milled samples) can be removed using extremely low energy (100eV - 1000eV) ion milling. Even at Ion energies of 500eV, you can operate at 20uA with a sputtering speed of 2.5 microns per hour. This new technolo= gy provides a means to reduce or eliminate amorphous damage without sacrificing milling rates. Obviously, purchasing a new system may not be= a feasible solution for you, but it is important to know that the technolog= y is available. =
Very useful information on Low Energy Ion Milling can be found in the paper: "Low Angle and Low Energy Ion Beam Etching for TEM Sample Preparation" by Arpad Barna. Proceedings of the Multinational Congress o= n Electron Microscopy Portoroz, Slovenia October 5-8, 1997. =
As I understand that this paper may not be easily accessible to many of you, I can make copies available to you at no charge if you send me your name and address. I also have many other papers on TEM sample preparatio= n which you might find of interest. If you have an interest, I can send yo= u a listing of the available papers.
Two other suggestions for your preparation may be to use the MicroCleave Technique which would eliminate ion milling altogether. The best people = to talk to about this technique are John McCaffrey (john.mccaffrey-at-nrc.ca) a= nd Scott Walck (walck-at-ppg.com).
The other suggestion is to minimize ion milling as much as much possible = by Tripod Polishing your samples into a wedge. You can produce a very thin sample which will require very little if any ion milling. The best conta= ct for this is Ron Anderson at IBM Analytical Services (anderron-at-us.ibm.com)=
The Pathology EM lab here still uses MT1s and MT2s to do thicks and thins. I thought I once saw an upgrade system for MT2s that would upgrade the knife stage to caliper movement and back lighting, and the specimen arc to caliper movement. Does anyone know if such a thing exists? I know the economics of upgrading an MT is a waste of money but I seriously don't think they would spend the additional money for even a used digital system. They are looking for a new tech and I will have to teach them how to section. My Ultracut and MT7 are older but so much nicer to teach with. If the new employee at least had smooth knife and specimen adjustment capabilities it would make learning so much easier. Thanks in advance.
Are you working on polycrystalline Solar Cells? That's what I was doing when I was involved with this material, trying to analyze it with a TEM. We usually had it on thick glass, approx. 3mm thick, with a metal contact between the CIS and the glass.
Let's see if I remember how we prepared that (It's been a few years):
The problem always was the glass, as you suggest in your email. We tried to transfer the CIS to Si before attempting to prepare it for TEM. The way we did that was by mechanical thinnig of the glass as far as possible. The sample was then glued onto Si with the CIS side down. After that, we dissolved the glass in HF. This left us with a much more manageable CIS on Si.
For cross sections, two, sometimes more, of these pieces (thinned) were then glued together and cross sections were prepared. To prevent them from falling apart, the TEM samples were stabilized with a small steel ring (Kim Jones from NREL had a Poster about this stabilizing technique at one of the MSA conferences, I believe). Further preparation then proceeded with mechanical dimpling and ion milling. I always used LN2 ion milling. Without this, artifacts would form, mostly In-rich "droplets". We were able to prepare beautiful samples with this technique, even with a metal backside contact. A 300 KV TEM may have helped, too. We could even do atomic resolution on these samples. That, however, may depend on your sample. if you have very small grains, they may fall out before the sample is done.
For plan views, I cut the TEM sample out of the Si, and thined it mechanically. It was then also stabilized with a steel ring. I then etched a hole into the Si from the backside. That way I was able to prepare free standing CIS films, which could then be ion milled. Perhaps that only worked because I had a metal contact underneath the CIS. It was a bit tricky, because the films sometimes would rip and curl.
I was told by the crystal growers that CIS is not soluble in HF and the comparison samples we prepared this way and the more traditional way never showed any differences.
Since I have not been working with this material for a few years, you should perhaps try to get in touch with Kim Jones at NREL. If you want to, I can get you his email. Please contact me through email.
Hope this helps.
Michael
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 phone: (888) FIND SIS fax: (303) 234-9271 email: info-at-soft-imaging.com
} ---------- } From: Albert Romano-Rodriguez[SMTP:romano-at-el.ub.es] } Sent: Wednesday, November 18, 1998 8:56 AM } To: Microscopy-at-sparc5.microscopy.com } Subject: TEM: Preparation of CuInS2 on glass } } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
The preparation of a concoidal "microchip" of your coating on a glass substrate provides a very thin edged sample suitable by itself as a plan view TEM specimen and it is also an excellent sample for ultramicrotomy to prepare a TEM cross-section. When this thin edged sample is broken to produce a thin and pointed sample it should be treated with an adhesion promoter, embedded in epoxy, cured and then sectioned using a diamond knife. CIS on glass should be a relatively easy material to section by ultramicrotomy.
Phil Swab Advanced Coatings Division/ART
} -----Original Message----- } From: Albert Romano-Rodriguez [SMTP:romano-at-el.ub.es] } Sent: Wednesday, November 18, 1998 10:56 AM } To: Microscopy-at-Sparc5.Microscopy.Com } Subject: TEM: Preparation of CuInS2 on glass } } ---------------------------------------------------------------------- } -- } The Microscopy ListServer -- Sponsor: The Microscopy Society of } America } To Subscribe/Unsubscribe -- Send Email to } ListServer-at-MSA.Microscopy.Com } On-Line Help } http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } ---------------------------------------------------------------------- } -. } } } Dear colleagues, } } We are working on the TEM characterisation of CuInS2 (usually } called CIS) on glass substrates. We are preparing TEM samples out of } these materials (both plan view and cross-section) and we are } encountering problems with the preparation procedure, which are } strong amorphisation of the CIS layer in plan-view and strong damage } (not always amorphisation) in cross-section. } } The preparation procedure we use is the standard preparation } method we use for Si-based materials: For cross-section, we cut } stripes out of the samples, glue them together. Next flat grinding, } dimpling and finally ion milling are used. For plan-views, a piece of } sample is ultrasonic cut and the preparation continues with the flat } grinding, ... and ion milling only from the backside. } } We believe that the problems we have are strongly related to the } glass substrate and to the charging of the glass during ion milling, } resulting in an overheating of the sample. This leads to extremely } poor cross-sections and to even worst plan-view samples (however we } still have some good results from few of these samples). It might be } also that the CIS layer is strongly beam sensitive (any experience } with it?). } } Any help in trying to circumvent these problems will be } extremely helpful and experience in preparing samples of the type } glass substrate-thin layer would also be strongly appreciated. } } By the way our ion milling machines can work at } liquid nitrogen temperature in order to minimise sample heating, if } this helps. } } Thank you in advance for your answers. } } Albert Romano-Rodriguez } Dept. of Electronics } Faculty of Physics } University of Barcelona } c/ Marti i Franques, 1 } E-08028 BARCELONA } Spain } tel: +34-93-402 11 47 } FAX: +34-93-402 11 48 } e-mail: romano-at-el.ub.es }
You may be losing some of the sulfur out of your sample when you ion mill. It is very easy to overheat the glass in the ion mill. You may also be having problems with In. See Cullis and Chew's paper on reactive iodine milling of In containing semiconductors in the MRS vol 115, 1988. You don't say what ion mill that you are using or the conditions. Someone has already suggested tripod polishing and that is a definite possibility, but I don't know how the CuInS2 reacts with water. (Try smelling it while you grind it on a 600 grit sheet of SiC with water as a lubricant. You should also smell the ion mill immediately after ion milling finishes.) Low angle milling might help you tremendously and it would avoid preferential sputtering effects and heating effects. I have not had problems with charging of glass samples but I mill at angles of less than 12 degrees and do quite a bit of it around 5-6 degrees at 5kV.
You could use the small angle cleavage technique with the glass if your films are adherent. The following steps are primarily for XTEM samples but if you read our paper that John McCaffrey sent to you, you will see that you can also pick out some samples that are suitable for plan view.
You may have problems with both water and heating your samples, but I don't know for sure. You can use SACT without water and without heating. Use a superglue for mounting instead of low temperature wax, but you will need to wait very long times to get the samples off of stubs with acetone to avoid heat. Use a viscous slow setting epoxy (super strong types that have about a 1hour working times) for mounting the samples on the grids. These take about a day to cure and again avoid heating the sample. This is explained in our paper. (MRS vol 480, 1997)
Modified steps for glass: 1. Grind the samples to about 75-80 um. on the backside 2. Assuming that you have rectangular samples, use a very coarse SiC paper (180 or 240 grit) and grind in straight lines parallel to the long axis of the sample. This will put your preferred crack directions into the glass, much like semiconductors have a preferred cleavage direction. When you flip the sample over, you will be able to see these scribe lines if your film is thin enough. 3. While the sample is still mounted, scribe the fracture lines at the 15 degree angle on the back side of the sample relative to these scratches, i.e. relative to the long axis of the sample. The distance between lines should be about 5mm. 4. Demount the sample, and break into the strips defined by the scribe lines. 5. Pick the strips of samples up and place on a post-it strip taped to a flat metal block with the sticky side of the post-it up. 6. Use your diamond miniscribe to "cleave" the sample along the scratches that were put there by the grinding with coarse SiC paper. 7. Place the pieces that you "cleave" that are sharp and possible good samples in the sticky corner of your post-it paper for examination, selection, and mounting on the girds for later.
Note: You can use No. 0 or 1 glass cover slips that are pre-ground to suitable thickness for your depositions. Then you can prepare the samples much faster.
South Bay Technology sells the "MicroCleave Kit" that John McCaffrey and myself had a hand in helping them put together. It has all the components that you need to get started doing the technique. With the kit, you really only need a good stereomicroscope.
I have no financial interests with SBT other than a customer, but I do have good friendships with the people that work there.
-Scott
Scott D. Walck, Ph.D. PPG Industries, Inc. Glass Technology Center Guys Run Rd. (packages) P.O. Box 11472 (letters) Pittsburgh, PA 15238-0472
Walck-at-PPG.com
(412) 820-8651 (office) (412) 820-8161 (fax)
"The opinions expressed are those of Scott D. Walck and not of PPG Industries, Inc. nor of any PPG-associated companies."
---------- } From: Albert Romano-Rodriguez To: Microscopy-at-Sparc5.Microscopy.Com -----------------------------------------------------------------------.
Dear colleagues,
We are working on the TEM characterisation of CuInS2 (usually called CIS) on glass substrates. We are preparing TEM samples out of these materials (both plan view and cross-section) and we are encountering problems with the preparation procedure, which are strong amorphisation of the CIS layer in plan-view and strong damage (not always amorphisation) in cross-section.
The preparation procedure we use is the standard preparation method we use for Si-based materials: For cross-section, we cut stripes out of the samples, glue them together. Next flat grinding, dimpling and finally ion milling are used. For plan-views, a piece of sample is ultrasonic cut and the preparation continues with the flat grinding, ... and ion milling only from the backside.
We believe that the problems we have are strongly related to the glass substrate and to the charging of the glass during ion milling, resulting in an overheating of the sample. This leads to extremely poor cross-sections and to even worst plan-view samples (however we still have some good results from few of these samples). It might be also that the CIS layer is strongly beam sensitive (any experience with it?).
Any help in trying to circumvent these problems will be extremely helpful and experience in preparing samples of the type glass substrate-thin layer would also be strongly appreciated.
By the way our ion milling machines can work at liquid nitrogen temperature in order to minimise sample heating, if this helps.
Thank you in advance for your answers.
Albert Romano-Rodriguez Dept. of Electronics Faculty of Physics University of Barcelona c/ Marti i Franques, 1 E-08028 BARCELONA Spain tel: +34-93-402 11 47 FAX: +34-93-402 11 48 e-mail: romano-at-el.ub.es
On Wed, 18 Nov 1998, valdemar wrote: } Dear Colleagues, } Could you kindly inform me if there is a forum similar to MSA for Raman Spectroscopy or micro-Raman Spectroscopy? } Berta } berta_m_t-at-hotmail.com
I established and maintain at Williams College a listserv called irusers-l. The listserv is subscribed to by a small number (about 68) scientists around the world who use FT-IR to analyze historic and artistic works. Some of the members, including myself, are interested in the complementary nature of raman, and would benefit from discussion of this technique, in addition to IR.
We're a quiet group, I suppose, but the low volume of discussion on the listserv has caused me to consider opening membership to IR spectroscopists (and possibly Raman spectroscopists) who do not work specifically with historic and artistic works.
My question is, are there members of this listserv who might be interested in participating in a listserv for IR spectroscopy and microspectroscopy or Raman?
James Martin Director of Analytical Services and Research Williamstown Art Conservation Center
Hello photon fans, I need an close loop, contactless system that can check & correct the working distance between objective & glass plate to within 1u. I would appreciate any feed back this group can provide. Please contact me directly if your company has markets this type of product.
I'd welcome feedback on an observation I made this afternoon, while examining a sample from an architectural finish, which included shellac, using epi-fluorescence illumination (details later).
The sample was dispersed in Cargille meltmount 1.662 between a glass slide and cover glass. When illuminated with wide-band ultraviolet illumination (HBO 100 Hg burner and UPlan fluorite objectives) gas began to evolve from the sample, displacing the surrounding mounting medium and leaving small voids within the coating sample. The rate of gas evolution increased with an increase in objective magnification/numerical aperture, and ceased when the vertical illuminator shutter was closed.
I have noted expansion of air pockets in samples before, and have witnessed coating layers melt in a cross-section sample before (both under epi-fluorescence illumination), but have never before witnessed uv-induced gas evolution from a sample. My first thought was a gaseous decomposition product from some unidentified component in the coating.
This is my curiosity of the day, and I'd appreciate hearing from anyone who has observed a similar behavior. Thanks.
James Martin Director of Analytical Services and Research Williamstown Art Conservation Center
Dear All, We are recently upgrading our Jeol JXA-8600 microprobe by a new ultra-thin window EDS detector. Does anyone know any solution to prevent or reduce a window contamination in that king of machine? How to improve vacuum in this, rather big, and contaminated microprobe chamber? Is that possible to clean this kind of detector window? Thank you for any advice and solution.
Vitaly Gutkin Electron Probe Laboratory
****************************************** ELECTRON PROBE LABORATORY Hebrew University of Jerusalem Institute of Earth Sciences Givat Ram, Jerusalem 91904 ISRAEL
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Hildy, Have you tried pre-embedding ICC? Fix your tissue with a high PAF-low Glut fixation and then vibratome to produce 30-50=B5m sections. Th= en process these for ICC using very small gold or PAP-DAB for final label.=20 Silver intensify the gold or DAB so it will be visible at both LM and EM levels. After ICC, you can osmicate and embed in any resin. We used to do = this with x-sections of rat brain and other tissues. We could check the sections by light microscopy and then embed those showing label. I would e= mbed between plastic coverslips which were weighted down with metal nuts so the tissue sections were very flat. Then I could cut around the edges of the cover slips, pop one off, and place gelatin capsules (with the solid end cut off) over the area of interest. A drop of resin would be put in the capsule and allowed to partially polymerize before filling the capsule and completing polymerization.
This had the advantage of maximum antibody penetration, permits osmium fixation so you can see the membranes, lets you examine tissue after=
labeling with both light and EM and select areas of maximum interest, and permits serial sectioning if necessary.=20 =20 Pleae contact for more details if desired. Debby -----------=20 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 =20 1057 Whistler Building West Lafayette, IN 47907-1057
Thanks to all who replied so promptly concerning the transfer of Data from a PC to A TN5500. I have one of two choices; Remove my printer and use that port, or try using a $MI string from library 55 (flextran) to accept data from TN Port 1. I may use the second option for a few reasons. I will be able to simply run a flextran program which transfers input to the PC, which then can control the TN5500 which controls the Cameca MBX Probe( hehe i think i need a flowchart to demonstrate this :) )
Those of you who asked how to recieve data from the TN (sending it to the PC), there are several options. To recieve text based data, i use a command } +DV 1. this changes my output from the printer to the serial port. I use hyperterminal to recieve the data on the PC side. As for images and such, I have two options. VISTA, a program supplied with our probe, is supposed to be an imaging program with outputs to the PC (I have never used this route though). We also purchased a GW electronics image capture board (slow scan analog to digital). I understand this board and the accompanying software is quite expensive.
Again thank you for your help Ted Claypool Engineer Scientist / EPMA RJ Lee Group
{!doctype html public "-//w3c//dtd html 4.0 transitional//en"} {html} Thanks to all who replied so promptly concerning the transfer of Data from a PC to A TN5500. {br} I have one of two choices; Remove my printer and use that port, or try using a $MI string from library 55 (flextran) to accept data from TN Port 1. I may use the second option for a few reasons. I will be able to simply run a flextran program which transfers input to the PC, which then can control the TN5500 which controls the Cameca MBX Probe( hehe i think i need a flowchart to demonstrate this :) ) {p} Those of you who asked how to recieve data from the TN (sending it to the PC), there are several options. To recieve text based data, i use a command } +DV 1. this changes my output from the printer to the serial port. I use hyperterminal to recieve the data on the PC side. {br} As for images and such, I have two options. VISTA, a program supplied with our probe, is supposed to be an imaging program with outputs to the PC (I have never used this route though). We also purchased a GW electronics image capture board (slow scan analog to digital). I understand this board and the accompanying software is quite expensive. {p} Again thank you for your help {br} Ted Claypool {br} Engineer Scientist / EPMA {br} {a href="http://www.rjlg.com"} RJ Lee Group {/a} {br} {br} {/html}
If I were you I would look at the Sem-Clean system from XEI Scientific. http://www.msa.microscopy.com/sm/xei
Their system bleeds nitrogen through the system to carry contaminants out of microscope, and it might be just the thing to clean up your probe.
best regards mark
Mark W. Lund, PhD VP Engineering } } 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
"This is a YOUNG business...How can I tell you what YOUR job is when I don't know what MINE is?" --Pogo
Dear All, We are recently upgrading our Jeol JXA-8600 microprobe by a new ultra-thin window EDS detector. Does anyone know any solution to prevent or reduce a window contamination in that king of machine? How to improve vacuum in this, rather big, and contaminated microprobe chamber? Is that possible to clean this kind of detector window? Thank you for any advice and solution.
Vitaly Gutkin Electron Probe Laboratory
****************************************** ELECTRON PROBE LABORATORY Hebrew University of Jerusalem Institute of Earth Sciences Givat Ram, Jerusalem 91904 ISRAEL
I had a chance to look at the cells I processed using HMDS and here's what I saw... They looked fine. I only went up to about 2,000X but they didn't look any different from the two samples that I ran up in the CPD. I used the slower infiltration of HMDS:
3:1 ETOH : HMDS 15 min. 1:1 E : H 15 min. 1:3 E : H 15 min. 100% HMDS 3 X 15 min.
Then I just put the tops to the dishes slightly askew and then let nature take it's course. I let them evaporate overnight in the hood. Then I put the coverslips onto stubs and sputter coated.
The one bad thing that happened was that some of the cells were grown in chamber slides and the HMDS ended up degrading the silicon seal enough for the stuff to leak out. The cells didn't look like they were too upset by this, though.
Whether this works for all cell lines, I don't know but I did this on loosely adherent tumor cells and lymphocytes.
Thanks to all who gave me suggestions as to what to try. Y'all are a truly great resource and I get lots of useful tips from this list.
Paula :-)
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
Hello collegues; Is there a forum (like this Microscopy one) for the X-ray phototelectron spectroscopy, or Auger, SIMS, ISS, etc.? If so, please let me know. Thanks. Sincerely yours Ram Srinivasan University of Kentucky
} ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } } Responding to the message of {3651F0D1.6C4F-at-botanica.ufrgs.br} } from Rinaldo Pires dos Santos {rinaldop-at-botanica.ufrgs.br} : } } } } ------------------------------------------------------------------------ } } The Microscopy ListServer -- Sponsor: The Microscopy Society of America } } To Subscribe/Unsubscribe -- Send Email to ListServer-at-MSA.Microscopy.Com } } On-Line Help http://www.msa.microscopy.com/MicroscopyListserver/FAQ.html } } -----------------------------------------------------------------------. } } } } } } Dear colleagues } } } } How to improve the contrast in plant tissue included in Spurr resin? The } } double contrast with Uranyl acetate in ethanol solution and Lead citrate } } is not good for me. There is some special procedure for to solve this } } problem? With the ultra-low viscosity resin, the problem is worst (lower } } contrast than the Spurr resin). } } Thank's in advance. } } } } M.Sc. Rinaldo Pires dos Santos } } Dept. of Botany - UFRGS } } Porto Alegre - RS - Brazil } } What kind of plant tissues? } } 1. Try a 50:50 mixture of Spurrs/Embed812 resins. Mix up resins seperately } without their accelerators (DMAE & DMP-30). You can store them in the freezer. } Then mix 50:50 just before use, mix in required amounts of the two accelerators, } infiltrate and cure. You should get the better sectioning and staining } properties of Embed812 plus the low viscosity of Spurrs. } } 2. Switch to Embed 812 100% and extend infiltration times. } } 3. What lead stain are you using? In my experience, Reynold's lead citrate } doesn't seem to work too well with Spurrs. I use Sato's triple lead stain, } preceeded by 3% UA, and get very good staining on plant and bacterial samples } embedded in Embed812. } } Good luck, } } Gib } } } } 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 } } } Dear Gib,
You got a problem! Please remember the following: Perfect infiltration (in your case the low viscosity) nixes contrast and immunocytochemistry attempts. You will not get perfect infiltration with epoxides, but then there are other problems. So, first, try this. 1. Use 1% Tannic Acid in cac buffer ,pH6.9, after glutaraldehyde fixation
2. Leave the prep in osmium overnight in the refrigerator.
3. Do an enbloc UA (3% maleate buffer, not phosphate or cac) for 90 minutes in the refrigerator
4. Add p-phenylenediamine to your 70% alcohol mixture - try 1% for 15min
5. Then look at some of the sections before staining in the TEM. Do not attempt to poststain those sections. See what you get.
6. Poststain with both UA and Reynolds. Reynolds yields more gray tones than Sato's.
7. If this does not work, then you must go to a very liquid epoxide, perhaps even a formulation with a dilutent in it.
Let me know if the above works.
Bye,
Hildy
P.S. Cut thicker sections. Use a smaller aperture! "Overexpose" (set up the density on your TEM) your negatives. Print on #1 and # 2 paper only. If you have to print on #3, your negative is not dense enough and you are loosing information. You would be amazed how diddling with the scope settings help! We drive this to extremes in cases with immunocytochem where we cannot use osmium UA, etc.
Note: The above sequence of TA, Osmium, UA, Ppd, UA, Pb, is a "cascade of mordanting agents" One acts as a mordant for the next all along.
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A single crystal of this material was jet electropolished with good success. A similar electrolyte may work for the CuInS2 material Mr. Rodriguez has. A south Bay 550 B single jet polisher was used with the following electrolyte: 60 ml. perchloric acid 460 ml. ethyl alcohol 280 ml. n-butyl alcohol 100 ml. butyl cellosolve 150 ml. acetic acid
Conditions: -20 degrees centigrade, 80 volts, 12 mA. current on a 1.5 mm. diameter exposed area. (One surface at a time). Lacquer protected the polished initial dimple during thinninging to automatic optical termination of the process. Some residue specks were seen on the foil, but more tests with the proportion of the ingredients might reduce that problem.
id {W04KZ8CD} ; Thu, 19 Nov 1998 16:47:38 -0500 Message-ID: {2D0F795D02A5D1118E7700A0C99B3AA23CC5A4-at-CHD64SMAIL01} To: microscopy-at-Sparc5.Microscopy.Com
*if replying please reply to this sender as he is "offlist".*
Hello Microscopy Experts,
At the suggestion of one of your colleagues, I would like to pose my inquiry to your list. I am not a subscriber, so I am sending this "off list" (hope you don't mind). If you wish to reply please reply to my email address {tim_wallace-at-doh.state.fl.us}.
Question 1. Are reference microphotographs available for natural fibers (cotton or cellulose for example)? I am trying to find a reference or photographic atlas to help me when looking in a light compound microscope at house dust samples. Can anyone suggest a web-site or a free/low cost resource with reference photographs or images that would be useful?
Question 2. Is there a lab in the United States that microscopically examines household dust, characterizing and identifying the contents (fibers, particles, hairs, globs etc.)?
I am Environmental Specialist working for a Local Health Department investigating a health complaint/concern from a resident. The resident is worried that the excessive dust problem, may pose a threat to her health. She really wants to know what is in the dust and where is it coming from. Any information would be appreciated. Thanks and best regards,
Tim tim_wallace-at-doh.state.fl.us
________________________________________________ Tim Wallace Environmental Specialist II Volusia County Health Department Environmental Health Division Indoor Air Assistance Lead Monitoring Programs 501 S. Clyde Morris Blvd. Daytona Beach, Florida 32114 USA phone: 904.947.3484 fax: 904.947.3485 http://www.state.fl.us/cf_web/D12/cphu/ehprgms/iaq.html +++++++++++++++++++++++++++++++++++++
XEI Scientific does make a cleaning system that can do what you need. I make a SEM-CLEAN nitrogen purge system for a JEOL 8600 probe, and I would be pleased to build another for you. Give me your mailing address and I will send you complete information.
Ronald Vane XEI Scientific 3124 Wessex Way Redwood City, CA 94061 (415) 369-0133
Vitaly Gutkin wrote: } } Dear All, } We are recently upgrading our Jeol JXA-8600 microprobe by a new } ultra-thin window EDS detector. } Does anyone know any solution to prevent or reduce a window } contamination in that king of machine? } How to improve vacuum in this, rather big, and contaminated microprobe } chamber? } Is that possible to clean this kind of detector window? } Thank you for any advice and solution. } } Vitaly Gutkin } Electron Probe Laboratory } } ****************************************** } ELECTRON PROBE LABORATORY } Hebrew University of Jerusalem } Institute of Earth Sciences } Givat Ram, Jerusalem 91904 ISRAEL } } VITALY GUTKIN } } Phone: 972-2-6585897 Fax: 972-2-5662581 } mailto:vit-at-cc.huji.ac.il } http://earth.es.huji.ac.il/e-prob/prob.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 } -----------------------------------------------------------------------. } } } } Dear Hildy } } I have had good results with neural preservation-- fixing with 4% } } formaldehyde (EM grade, no methanol) +.2% glutaraldehyde in PBS. } } Dehydration was partial and in a methanol series with progressive } } lowering of temperature, } } infiltrated in LR white at -20C, and polymerizing at 55C. The series of } } methanol was 60% for 15' at 4C, 70% methanol for 60' at -20C, and 80% } } methanol for 60' at -20C, then 3 changes of LR white for 60' at -20C. Hope } } this will help. This method has given good antigenicity with a wide } } variety of tissues. If you need further info, please feel free to contact } } me. } } Marge } } } } Margaret Springett } } e-mail hukee.margaret-at-mayo.edu } } IEM Specialist at Mayo Foundation } } 1426 Guggenheim } } Rochester, Mn. 55905 } } } } Margaret Springett } e-mail hukee.margaret-at-mayo.edu } IEM Specialist at Mayo Foundation } 1426 Guggenheim } Rochester, Mn. 55905 } } } } Hi,
The above is an excellent protocol. We have found it to visualize antigens which absolutely will not allow themselves to be detected any other way. We are getting great label. Our problem is that we can only use 0.25% GA (I hate this antigen) and the structure which we need to show up (vesicles) do not seem to preserve enough for us to call them vesicles in a publication. I recommend your protocol highly to anyone who has trouble with epoxides in immuno. (Our preservation in expoxides was quite good even without osmium, but vesicles tend to be ephemeral anyway, and the poor preservative ability of the LRs seems to have put them over the edge). Thanks, Hildy
Vitaly Gutkin wrote: {snip} } How to improve vacuum in this, rather big, and contaminated microprobe } chamber? } Is that possible to clean this kind of detector window? } Thank you for any advice and solution. } } Vitaly Gutkin } Electron Probe Laboratory } } ****************************************** } ELECTRON PROBE LABORATORY } Hebrew University of Jerusalem } Institute of Earth Sciences } Givat Ram, Jerusalem 91904 ISRAEL } } VITALY GUTKIN } } Phone: 972-2-6585897 Fax: 972-2-5662581 } mailto:vit-at-cc.huji.ac.il } http://earth.es.huji.ac.il/e-prob/prob.html } ******************************************
First, of course, clean the chamber well.... Perhaps water/detergent, alcohol, acetone, etc... will depend on cantaminants.
The windows can be cleaned by carefully dripping solvent across the window. There has been a thread about this recently on the Microscopy Society of America Listserver.
Keeping it clean:
Not familiar with specifics of your JEOL, but have been running a Be/UTW/Open Window EDS detector for 15 years and never had an oil problem. Only a few times have I had to warm the detector to clear the (cryo pumped) crystal. Most of this problem I attribut to P-10 gas (argon/methane) from a less-than-perfect WDS thin window leak. The key may not be cheap, depending on your configuration. My SEM is an ETEC which has staged vacuum pumping. The chamber is rough pumped to 100 microns. At this point vacuum valving places a Magnetic levitation bearing turbo pump ahead of the rough pump. ..Very clean vacuum. The only improvement would be to use an oil-less rough pump, but I haven't had the funding.
Regards, Woody
--------------------------- -- ------------------ de Woody, WB4QXE -------------------
- Work: SEM/EDS/WDS - Materials Research & Failure Analysis also, electronics and instrumentation.
- Home: Ham radio "homebrewing", computers , shade tree mechanic.
- www site: Scanning Electron images and Ham Radio Homebrewing stuff. http://www.geocities.com/capecanaveral/3722
The only forum I know is for SIMS. The web address is listed below. You can also subscribe to a list server from the site. http://www.simsworkshop.org/default.nclk
Hope this helps, Ed Hirsch
At 11:22 AM 11/19/98 -0800, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
I am still working on my project to teach histology to highschool students. People on this list have been very helpful in getting me a microtome and prepared slides. I am now working on the chemicals & supplies I need. Going over an on-line catalog I discovered that there are much safer substitutes for the chemicals used in my 1962 edition of "Animal Tissue Techniques" by Humason. However, it was not clear to me what safer solvent is used to substitute for toluene in the paraffin infiltration stage. Can anyone suggest an up to date reference book which discusses the advantages and disadvantages of the new substitutes. It appears that a lot has changed since I worked in a medical lab 30 years ago. Thanks.
The best source for particle information is from McCrone Accessories. It is called the Particle Atlas and may be available through a local university science library. Commerically, it is available on CD Rom, for about $1100. It is a really great resource (we've used it with several clients and classes). The McCrone Institute is also a good lab for this type of thing. Closer to your area is a high quality spin-off, run by Skip Palenik. The contact numbers for both MAC and the Institute are below. I don't have current info for Skip but imagine that you can find him on the Web.
} Dear All, } We are recently upgrading our Jeol JXA-8600 microprobe by a new } ultra-thin window EDS detector. } Does anyone know any solution to prevent or reduce a window } contamination in that king of machine? } How to improve vacuum in this, rather big, and contaminated microprobe } chamber? } Is that possible to clean this kind of detector window? } Thank you for any advice and solution. } } Vitaly Gutkin } Electron Probe Laboratory } } } ****************************************** } ELECTRON PROBE LABORATORY } Hebrew University of Jerusalem } Institute of Earth Sciences } Givat Ram, Jerusalem 91904 ISRAEL } } VITALY GUTKIN } } Phone: 972-2-6585897 Fax: 972-2-5662581 } mailto:vit-at-cc.huji.ac.il } http://earth.es.huji.ac.il/e-prob/prob.html } ****************************************** } Dear Vitaly! The radical solution is to replace diffusion pump oil by Santovac-5. I made it some years ago on the predecessor of your microanalyzer - JEOL Superprobe JXA-733. In result the vacuum was improved everywhere on the order. Contamination of sample under beam have decreased considerably (analysis of light elements has become more accurate). Besides I have become to clean the column 1 time per 4 years (before it was 1 time per 3 months). Precautions: 1. Santovac-5 has more viscosity therefore probably it will be necessary to increase slightly the power of diffusion pump heater, for example by autotransformer. 2. To notice effect it is necessary to clean well the column and vacuum system from old oil (most labour-consuming procedure). I would like to advise also to install a trap for forepump oil vapours and the liquid nitrogen trap for diffusion pump and plate above sample holder. Though from conference it is known all EDS manufacturers make windows in one place, it seems better to request of the manufacturer for recommendations on window cleaning. For example Oxford Instruments has this information. Regards.
Victor Sidorenko, ANTRON Co. Ltd., scientific service, Moscow, Russia antron-at-space.ru
The only forum I know is for SIMS. The web address is listed below. You can also subscribe to the SIMS list server from the site. http://www.simsworkshop.org/default.nclk
Hope this helps, Ed Hirsch
At 11:22 AM 11/19/98 -0800, you wrote: } ------------------------------------------------------------------------ } The Microscopy ListServer -- Sponsor: The Microscopy Society of America
} } In regards Beth Richardson's search for a new blade for the Dahle paper cutter. You can get one. I have had my Dahle for over 10 years and replaced the cutting blade about a year ago. I called my local photography supply house and they ordered a new blade for me. It was easy to put in and I've been using it ever since. It's cheaper than buying a new cutter. God luck. } JoAnn Buchanan Molecular and Cellular Physiology Stanford University School of Medicine Stanford, CA 94022
One of the best histology books out today is=20 Histotechnology, A Self-Instructional Text=20 by Freida L. Carson
ISBN 0-89189-306-7
You can be ordered the book through the American=20 Society of Clinical Pathologists
George Lawton Chief Electron Microscopist Microscopy and Imaging Center UT Southwestern Medical Center at Dallas Phone: 214-648-7291 eMail: George.Lawton-at-email.swmed.edu
Why is Hepes buffer not used in the primary fixation with glutaraldehyde for mammalian tissues instead of the phosphate and cacodylate buffers? Anyone know the biochemical reasons?
Apparently in my last posting concerning Low Energy Ion Milling, Tripod Polishing and the MicroCleave Technique my disclaimer and signature line did not appear. While it did appear on my copy of the message, for some reason it did not appear on the copy of the message that I saw on the Listserver. I am sure that the people on the list know that we produce these products, but I thought I should make it clear by reiterating my disclaimer here. So here goes:
NOTE: We do sell the IV3 Ion Milling System with LEGs (Low Energy Guns) a= s well as the MicroCleave Kit and the Tripod Polisher so I do have a vested=
For some of us, HEPES is the buffer used for primary fixation with aldehydes. I find it works fine. PIPES would be another good choice (it has a slightly lower pKa so would have greater buffering capacity that HEPES if both started at 7.4 and the solution had a tendency to acidify (like aldehyde fixatives do).
} Hi again, } } Why is Hepes buffer not used in the primary fixation with glutaraldehyde } for mammalian tissues instead of the phosphate and cacodylate buffers? } Anyone know the biochemical reasons? } } Thanks, } Hildy
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)
Skip Palenik is on the Web at palenik-at-aol.com and is an excellent resource. His Company Name is Microtrace in Elgin, IL Phone: 847-742-9909
I have no financial interest in this recommendation.
Tom Kremer Analytical Science & Technology Kimberly-Clark 920-721-4583 e-mail: tkremer-at-kcc.com
} ---------- } From: Barbara Foster[SMTP:mme-at-map.com] } Sent: Friday, November 20, 1998 9:26 AM } To: Tim_Wallace-at-doh.state.fl.us-at-sparc5.microscopy.com; } microscopy-at-sparc5.microscopy.com } Subject: Re: Microscopic House Dust Characterization & ID } } ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } Tim, } } The best source for particle information is from McCrone Accessories. It } is called the Particle Atlas and may be available through a local } university science library. Commerically, it is available on CD Rom, for } about $1100. It is a really great resource (we've used it with several } clients and classes). The McCrone Institute is also a good lab for this } type of thing. Closer to your area is a high quality spin-off, run by } Skip Palenik. The contact numbers for both MAC and the Institute are } below. I don't have current info for Skip but imagine that you can find } him on the Web. } } } McCrone Accessories: 800-622-8122 } McCrone Institute: 312-842-7100 } } Best of luck, } Barbara Foster } Consortium President } Microscopy/Microscopy Education ..Educating microscopists for greater } productivity. } } 125 Paridon Street Suite 102 Springfield, MA 01118 } PH: (413)746-6931 FX: (413)746-9311 email: mme-at-map.com } Visit our web site {http://www.MME-Microscopy.com/education} } ****************************************************** } MME is America's first national consortium dedicated to } customized on-site training in all areas of } microscopy, sample preparation, and image analysis. } } } } At 04:45 PM 11/19/98 -0500, } Tim_Wallace-at-doh.state.fl.us"-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 } } -----------------------------------------------------------------------. } } } } } } *if replying please reply to this sender as he is "offlist".* } } } } Hello Microscopy Experts, } } } } At the suggestion of one of your colleagues, I would like to pose my } inquiry } } to your list. I am not a subscriber, so I am sending this "off list" } (hope } } you don't mind). If you wish to reply please reply to my email address } } {tim_wallace-at-doh.state.fl.us}. } } } } Question 1. Are reference microphotographs available for natural fibers } } (cotton or cellulose for example)? I am trying to find a reference or } } photographic atlas to help me when looking in a light compound microscope } at } } house dust samples. Can anyone suggest a web-site or a free/low cost } } resource with reference photographs or images that would be useful? } } } } Question 2. Is there a lab in the United States that microscopically } } examines household dust, characterizing and identifying the contents } } (fibers, particles, hairs, globs etc.)? } } } } I am Environmental Specialist working for a Local Health Department } } investigating a health complaint/concern from a resident. The resident } is } } worried that the excessive dust problem, may pose a threat to her health. } } } She really wants to know what is in the dust and where is it coming from. } } } Any information would be appreciated. Thanks and best regards, } } } } Tim } } tim_wallace-at-doh.state.fl.us } } } } ________________________________________________ } } Tim Wallace } } Environmental Specialist II } } Volusia County Health Department } } Environmental Health Division } } Indoor Air Assistance Lead Monitoring Programs } } 501 S. Clyde Morris Blvd. Daytona Beach, Florida 32114 USA } } phone: 904.947.3484 fax: 904.947.3485 } } http://www.state.fl.us/cf_web/D12/cphu/ehprgms/iaq.html } } +++++++++++++++++++++++++++++++++++++ } } } } } } } } } } }
In a message dated 98-11-20 17:07:30 EST, hcrowley-at-du.edu writes:
{ { Does anyone know if enbloc uranyl acetate staining interferes with UV Polymerization of LR White or LR Gold at -20C? Has anyone done it? Thanks, } }
Hi Hildy,
I don't know about LR White and Gold, but en block stains can sure cause problems with Lowicryl K4M, which is also polymerized by UV. I would guess that you may encounter some problems, probably in the centers of the specimens.
Cheers, Bob **************************************** Robert (Bob) Chiovetti, Ph.D. Microimaging Technologies, Inc. Tucson, Arizona USA Tel. / Fax (520) 546-4986 rchiovetti-at-aol.com Manufacturers' Representatives Systems Integrators Analog & Digital Imaging Systems *****************************************
The following showed up on the Sci.Techniques.Microscopy Usenet Newsgroup. Someone on this list may be able to help him. If so, please respond directly to him at:
emans-at-bio1.rwth-aachen.de
Please don't respond to this message you are reading now, since I have only copied the text into this message.
Thanks.
Bob Chiovetti
***************************************
I keep seeing posts mentioning the McCrone Research Center on this list server. I have never seen it mentioned that they have a web page. So for those interested, they do. It is:
HILDEGARD CROWLEY wrote: } } Hi, } } Does anyone know if enbloc uranyl acetate staining interferes with UV } Polymerization of LR White or LR Gold at -20C? } Has anyone done it? } Thanks, } Hildy
"Use of uranyl acetate en bloc to improve tissue preservation and labeling for post-embedding immunoelectron microscopy" Erikson, P.A. et al. 1987. J. Elec. Micros. Tech. 5:303-314.
They stained en bloc with UA before embedding in LR White or Lowikryl K4M. Of course, the antigen in question might make a difference.
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 ***************************************************************
We are planning to rebuild the optical diffractometer we use to assess the quality of our TEM micrographs. Anyone have a favorite design or modification for an optical diffractometer (and perhaps an associated reference)? We are looking to build a vertical set up similar to the one in:
We'd also like to add a CCD camera with a high quality thermal printer.
Any recommendations?
Thanks,
- Dennis
------------------------------------------------------------------------------ Dennis C. Winkler, PhD. Phone: (301) 496-0131 Laboratory of Structural Biology Research Fax: (301) 480-7629 NIAMS, National Institutes of Health Email: Dennis_Winkler-at-nih.gov Bldg. 6, Room B2-26, MSC-2717 Bethesda, MD 20892-2717, U.S.A.
To Harris Reavin, The evolution of clearing solvents is interesting. Toluene (actually chloroform) works best, but is a no-no because of carcinogenicity and environmental (disposal) problems. Xylene has been the solvent of choice for both clearing and dewaxing, but again, health effects and disposal are major concerns. Limonene products came on the scene as the "magic" xylene substitute, boasting the fact(?) that they could be dumped down the drain. Well, in my experience they don't perform well, our local sewer authorities don't want them in the system and you can smell them out in the parking lot! Aliphatic hydrocarbons, such as certain products from Richard-Allen Scientific 800-522-7270 and ANATECH 800-ANATECH have a proven track record as xylene substitutes. Check with these companies, you might persuade them to send you some samples...and they both are very knowledgeable and helpful and offer all kinds of expert advice. In my opinion, you'll get good clearing with aliphatic hydrocarbons, but you will have to do your whole processing procedure under a fume hood. Remember gloves, goggles and apron...be safe! Good luck, Bob Santoianni Emory University Hospital Atlanta, GA robert_santoianni-at-emory.org
Grateful thanks for the helpful advises and comments to anybody, who response my questions about JXA-8600 microprobe cleaning.
Yours sincerely,
Vitaly Gutkin Electron Probe Laboratory
-- ****************************************** ELECTRON PROBE LABORATORY Hebrew University of Jerusalem Institute of Earth Sciences Givat Ram, Jerusalem 91904 ISRAEL
The E.M. Unit in Triniy College Dublin have a secondhand Emscope SP2000A Cryogenic preparation system for sale. This is a fully working system with a gas cooled stage, rather than the usual braid system, to give a rapid response to cooling and heating requirements. The asking price is =A35,000 sterling with the purchaser arranging carriage and insurance. =46or full details please contact me off list.
David John, Manager, Electron Microscope Unit, Trinity College, Dublin 2, Ireland. tel.no. (353) - 1 - 6081559 e-mail - djohn-at-mail.tcd.ie
Last week during the thread on CuInS2 on glass, someone from Raytheon somewhere in Texas sent me a personal email asking for a copy of the Small-Angle Cleavage Technique video tape. We routinely mail these out like a paper reprint, but I inadvertantly trashed his message and lost his mailing address. So, will that person from Raytheon (or anyone else who wants the video) please send me another message, and I'll send him a copy of the tape. My apologies to everyone else for filling up your email message box.
Cheers John
John P. McCaffrey Institute for Microstructural Sciences National Research Council of Canada M-50 Montreal Rd. Ottawa, Ontario K1A 0R6 Canada
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
RE} McCrone Res. Center on web 11/23/98 8:55 AM Thank you Azriel for the mention of the McCrone web site.
The "home" web address is:
http://www.mcri.org
} From there you may go to any of our pages and find out about us, our cour= ses, our expertise, mission, etc.
Thanks.
John D. Shane Director of Research McCrone Research Inst. 2820 South Michigan Ave. Chicago, IL 60616 312.842.7100
--------------------------------------
I keep seeing posts mentioning the McCrone Research Center on this list server. I have never seen it mentioned that they have a web page. So for those interested, they do. It is:
Does anyone know how the get information on durability and storage of B&W prints from the thermal printer. We use Seikosha VP-3500 printer, but we were not able to get any information from the manufacturer.
I am looking for a used color monitor for a Kevex Delta 1 EDX System. It is a model 38-DO5IMA-UU free standing monitor with 5 RCA plugs exiting the rear.
If anyone can help, please contact me at 303-689-2224 or e-mail franklin-at-idcomm.com...
Hi all. Yes this is real boring technical stuff but we hope it helps somebody. A while ago we asked if any one else was having problems with some of = the older ISI SEM's regularly blowing the transistors in the box on top = of the HT tank. We had quite a response from many people having the same = problem but not may solutions to the problem that seemed to work.
We think we have been able to identify the problem and also solve the = problem.
The cct inside the tank on the bias and HT side that has the 4700pF and = 2200pF caps on it, is to filter out the back emf from the high voltage = side of the tank. We found that these same caps go soft and so leak that = voltage back up into the driver transistors on top of the tank. This is = what causes them to blow. It is quite simple to monitor this. If you = measure the collectors of these transistors you should measure waveforms = with a peak voltage not exceeding 400v on any of the transistors. ( the = actual voltage will vary with bias settings, filament current and HV.) = Should you find that the voltage is close to this voltage or over, then = replace the relevant caps in the tank ( C20, C21, C4, C5 etc. 4700pF = 2.5Kv and 2200pF 4Kv and 0.1uF 630v) This will solve the problem.
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
Fellow microscopists, I am setting up a darkroom for my electron microscope laboratory recently and looking up a suitable enlarger. I would appreciate your suggestions and comments on what brand/ specifications I need for my TEM, SEM and optical microscopes. Thanks Ren-Jye
We have a Seikosha VP-3500 and I can give you some input.
These prints will degrade very rapidly if left out in the lab where they are exposed to fluorescent lights. You can see degradation beginning to occur in a week or so.
Best bet is to store them in a dark cabinet, this will help slow the degradation process. I have pictures that are several years old stored that way.
I wouldn't count on them for long term storage of critical information. we went to a relatively cheap digital image capture system (Snappy Image capture card on the NTSC outlet of our AMRAY 1830) and are well pleased with the results. We typically take both thermal prints and a digital image on all images.
John Giles Principal Materials Engineer Honeywell Space Systems
} Hi everyone,
} Does anyone know how the get information on durability } and storage of B&W prints from the thermal printer. We } use Seikosha VP-3500 printer, but we were not able to } get any information from the manufacturer.
On Mon, 23 Nov 1998, Craig Franklin wrote: } I am looking for a used color monitor for a Kevex Delta 1 EDX System. It is } a model 38-DO5IMA-UU free standing monitor with 5 RCA plugs exiting the } rear.
If those plugs are labelled R,G,B,H,V, it is probably a standard RGB monitor and there are many replacements from many manufacturers. If it is an older machine, the bandwidth is probably not great making replacement relatively easy. Surely, someone in the lab/department has a computer monitor with RGB inputs you could try out.
: On Mon, 23 Nov 1998, Craig Franklin wrote: : } I am looking for a used color monitor for a Kevex Delta 1 EDX System. It is : } a model 38-DO5IMA-UU free standing monitor with 5 RCA plugs exiting the : } rear. : : If those plugs are labelled R,G,B,H,V, it is probably a : standard RGB monitor and there are many replacements from : many manufacturers. If it is an older machine, the : bandwidth is probably not great making replacement : relatively easy. Surely, someone in the lab/department has : a computer monitor with RGB inputs you could try out. :
The monitor you have was most likely made by Electrohome.
A standard RGB monitor will work, but only if it can sync at low H sync rates. Our 8000 puts out an H sync of 17 Hz and a V sync of 57 Hz.
Carl
Carl Henderson Electron Microbeam Analysis Laboratory University of Michigan 2501 C.C. Little Bldg. Ann Arbor, MI 48109-1063 USA -------------------------------- Voice: (734) 936-1550 FAX: (734) 763-4690 E-mail: chender-at-umich.edu --------------------------------
Sorry for the delay in summarizing all the feedback ... I figured the best way would be just to cut-and-paste everyone's advise onto one email because I could miss certain points.
All advice has been a great help.
Thank you, Ad
+++++++++++
} From:Yasuo Ito
Hi, I got your message through microscopy list server. Yes, i think you have a tough assignment since sulphur L-edge(I presume that you are looking at these edges) is at about 165 eV. My guess is that (1) the thickness of your specimen may be too thick (2) interefered by any other edges, and/or (2) sulphur might be removed due to the electron-beam damage.
For (1), if the speciment is too thick, the plasmon loss peak becomes to big and swamp the sulphur peak. How thick is your specimen? I mean how thick in terms of inelastic mean free path. You can measure this by taking low loss spectrum of the are of interest. I guess, the thickness would be less than a half of that.
For (2)and (3), this depends on what is the matrix of your specimen.
For (3), you can monitor any change of the mass thickness by monitoring plasmon peak, as you may know. Or at least you may be observe by images. Have you seen any indication of the beam damage? Do you have EDS? you may be able to check by the EDS for the existence of sulphur in your specimen. If the beam damage is the case, you may have to reduce the dose into the area by reducing beam current or acquisition time.
So far this is what I can think of at hand. I hope these would give you some idea (I guess you may have already thought about these possibility, though.)
Please don't hesitate to contact me for any further questions. I have been dealing with beam sensitive materials for a while.
Cheers,
Yasuo
++++++++++++
} From:Larry Thomas
Advice is cheap, so I'm told, but here goes.
I would take not being able to see the sulfur edge as a bad sign. Maybe there is really no S in your sample. Are you sure the 'sulfur ' peak you saw by EDS wasn't actually a Mo peak or Pb peak from the sample or from a microscope artifact. It's not unusual to get a small Mo x-ray contribution from scattering off the condenser apertures in the TEM. You say the sulfur concentration in the sample "is not that much." How much is that? The detection limit in EELS might be as much as several percent. If the signal is being hidden by the huge background in EELS, the most sensitive way I know of to detect it is by using 2nd difference spectra. Difference collection modes remove background much more effectively than power law subtraction. There is an optimum collection angle in EELS, so you might try different collection apertures. If you can't detect the sulfur in difference spectra with enough counts for the detection limits you need, don't expect anything useful from power law correction, and especially not from mapping with very limited counts from individual analysis points.
I would also ask myself the question "If I'm getting easily detectable sulfur by EDS analysis, why bother mapping with EELS?"
Anyway, that's my two-cents worth.
Larry Thomas Washington State University
+++++++++++++++
} From:Gerd Duscher
I am working mostly with a VG HB501 with Gatan PEELS for several years. So in principle I have the same sensitivity for sulphure as you.
I think your experimental conditions are fine!
I had a similar problem with Silicon, which I solved using two different approaches.
First I would collect two spectra with good counting statistics in the sulphur and in the low loss region. Splice them and do the single scattering deconvolution using the LOG-ratio method in EL/P. This ensures that you get always the same background before the edge and you cancel out thickness effects.
Secondly, if you have regions without sulfur and without, you can use the spatial difference technique: author = {H. M\"ullejans and J. Bruley}, title = {Improvements in Detection Sensitivity by Spatial--Difference Electron Energy--Loss Spectroscopy at Interfaces in Ceramics}, journal = {Ultramicroscopy}, year = 1994, volume = 53, number = 4, pages = {351-360},
You need two spectra which are taken under the same conditions and at locations with the same thickness. Then you use the one without for an improved background subtraction.
I have just noticed your message about sulfur mapping. I wonder if you have seen the following publication?
Spatial distributions of sulfur-rich proteins in cornifying epithelia. Leapman, RD, Jarnik, M, Steven, AC. J Struct Biol 1997; 120: 168-179.
I shall be pleased to send you a reprint if you give me your mailing address.
...Richard Leapman
_____________________________ National Institues of Health Bioengineering & Physical Science Program, ORS Supramolecular Structure & Function Resource Bldg. 13, Rm. 3N17 Bethesda, MD 20892-5766 tel: (301) 496-2599 fax: (301) 496-6608 e-mail: leapman-at-helix.nih.gov
web reference: http://www.nih.gov/od/ors/beps/ssfr/ _____________________________
+++++++++++++++
} From:Charles A. Garber
This might be a dumb kind of comment, so forgive me if you think it is, but one more than a few instances, people have found Mo in their spectra coming from Mo apertures. So you might want to check and see if you are using Mo apertures and if yes, you might want to change them to apertures of some other composition (for example, Pt) and see if your Mo peaks are still there or not.
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: www.2spi.com ############################
++++++++++++++++++++
} From: Michal Jarnik
Ad,
we recently published an article in the Journal of Structural Biology (Leapman, R. D., Jarnik, M. and Steven, A. C. (1997). Spatial distributions of sulfur-rich proteins in cornifying epithelia. J. Struct. Biol., 120, 168-179). If you send me your mailing address I will be happy to get you a copy.
Regards,
Michal -- Michal Jarnik, Ph.D. Fox Chase Cancer Center Electron Microscopy Facility 7701 Burholme Ave. Philadelphia PA 19111 Tel. 215-728-5675 Fax 215-728-2412
++++++++++++++++++
} From: Haswell Malcolm
I believe that one of the other listers has already mentioned molybdenum peaks. This has been a chronic problem on our Hitachi H7000 because of the Mo in the movable objective apertures.
We have had problems with EDX, not EELS, but I would expect that the geometry may be worse in EELS because your detector might more directly "see" objective apertures and so increase the chance of picking up Mo. If your system will pick up higher energies of Mo it would certainly be worth doing a quick check.
Incidentally I received a lot of advice about our molybdenum problem some time ago and I can't remember if I thanked everyone - so thanks, just in case. Unfortunately all we ever managed to do was minimize the effect, unless we removed the aperture rod which produced its own problems.
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD Tyne and Wear UK
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk
++++++++++++++++++++++
} From:Laryy Thomas
Gerd's suggested method for removing background is technically the best although it involves a bit more work than difference spectrum collection.
You haven't said what S concentration you're actually looking for. What is it?
To look for hard-to-find sulfur in EELS, I'd find the S-containing area with EDS before trying EELS. In EDS or EELS, use an area collection mode such as selected-area diffraction rather than a focused probe mode to minimize beam-damaging the sample. A 50 nm sample thickness should be fine. The broad type of edge sulfur has at ~165 eV is easy to hide in background. At that energy loss, my experience is a power law gives very poor fits. If you have the Gatan DigiPEELS system, has anyone mentioned the correction for point spread in the detection system? It can be important.
As to the comment about preferring EELS mapping over EDS mapping because of much longer necessary acquisition times in EDS, I don't see why this should be so. Certainly you should optimize the acquisition conditions in EDS (you can use a short processing time and high beam current to optimize counting rates), but you have to do essentially the same thing in EELS anyway. The question to ask yourself is "What detection limit do I need for this job and therefore, what count rate do I need to get good statistics in a reasonable acquisition time?" Another question is "Why do I really need a map?" Mapping is pretty much a cosmetic operation anyway--good for show and tell or for impressing others, but not the way to detect elements at small concentrations. I'd use point or area analyses to get the necessary counting statistics.
You can get the header file for EL/P by using the program's facility for saving/reading files as text. If you're only converting a few files, use cut and paste.
Is Mac Fac a multivariate stat. analysis program? Where can I get it?
Larry Thomas
++++++++++++++++++++++
} From: Tony Garratt-Reed
I don't know what responses were made to Daraporn off the listserver, but there seems to be some confusion in what has been posted on the server.
Daraporn has seen a small peak at the S energy in his EDX spectra, but could not find S in the EELS spectrum. This could be for one of two reasons: 1) the S is not S at all, but Mo or Pb. In this case, with luck, he might expect to see the K-lines of Mo or the L-lines of Pb at higher energy, provided, of course, that there is enough present (the low-energy lines at the S energy are much more intense than the higher energy ones) Mo could certainly be coming from apertures, as Malcolm Haswell suggests. Pb could, conceivably, be coming from the x-ray shielding in the microscope, but do other CM20's have that problem? There is no obvious reason why Daraporn's should if others don't, unless they have made some modification to the stage area. It would seem unlikely that Mo or Pb were actually in the sample. EELS would be used to confirm the analysis because EELS is not susceptible to "hole count" effects. To use the Mo aperture as an example: the Mo x-ray signal is generated by stray radiation (scattered electrons, x-rays)hitting the large area of the mostly thick aperture material. Hence just a few electrons or x-rays can produce lots of Mo x-rays. In contrast, the few electrons that penetrate the aperture would have lost lots of energy and been scattered through large angles, so would not reach the EELS detector at all. There is no equivalent of the mechanism that generated the spurious x-ray signal.
Unfortunately, there is another possible reason for not seeing the S, even if it is really there. The sample could be too thick. The whole relationship of beam current, sample thickness, statistics and spatial resolution is very complex, both in EDX and EELS analysis. However, one point is probably worth mentioning: Comparisons of the detection sensitivities for EDX and EELS compare the results from the same samples. Typically, in practice samples suitable for EELS are thinner than those often used for EDX analysis. Hence, although theory might say that (at least up to Ca or so) the detection limits for EELS are better than those for EDX, this is only true for thin samples. In thicker samples, the EELSbackround rises and the peak becomes blurred out, resulting in poorer detection, while in EDX you just get more signal and hence better detectability.
The papers (from NIST/NBS) that illustrate the sensitivity of EELS also point out the need for extremely high incident beam currents, available only in a LaB6 microscope. I don't know enough about the CM20 FEG to translate from the settings that Daraporn used to actual beam characteristics. However, it is clear that the optimum beam settings for EELS are very different from those for EDX, and also the spatial resolution is significantly degraded. It could be that his microscope conditions are not close to optimum for EELS.
I don't know what is the approx. S-content ... they just told me "it's low" .. but with my parallel probe analysis with 100sec collection time, meter plate reading of 2.5 sec ... the maximum sulfur peak with EDS was ~1000 counts. Not sure what the meter plate is. It might be what I called the processing time in the EDS system. This is essentially the EDS pulse processor time constant. Short processing-times allow high count rates at reduced deadtime, although with some degradation in energy resolution. Long P-times give the best resolution but increase deadtime at a given counting rate. For mapping, you sacrifice energy resolution to allow the highest possible counting rate: set the signal processor for the shortest processing time. But when I switch to do the analysis in STEM mode, because the probe is small, the sulfur peak has to be collected with a longer time. Whatever mode you choose for analysis, you can optimize the counting rate by increasing the probe size and using a large condenser aperture to allow a large beam convergence angle. Nearly all TEMs allow you to change these parameters: you're not limited to the 'standard' microscope settings. You're also not limited to STEM mode. Did Tony Garrett-Read mention that you were using a CM20 FEG? I don't know which EDS system you're using, but most can give at least 30,000 counts./sec with less than 50% deadtime. Sure, you sacrifice spatial resolution, but you'll still get plenty for most jobs, including yours, even with a conventional TEM. To check the sulfur detection, I'd highly recommend spreading the probe to avoid damaging the sample. If you decide to proceed with mapping, first run a little side experiment comparing analyses with a focused probe and a spread one to see if the sulfur is stable. While with EELS, I only need 5 sec or something like that. A 5 second EELS analysis sounds as if you're not counting long enough to get the statistics for a reasonable detection limit.
I need the mapping mainly for the cosmetic purpose and because it's give us spatially resolved information, something like that. That's the term they used.
You mentioned about "point spread in the detection system" in DigiPEELS, what is that? No one mentioned about that. Is that in the newer version? DigiPEELS is Gatan's newest model PEELS detection hardware. If you don't have it, don't worry about it. Newer versions of the EL/P software have a spectrum sharpening function that corrects for the interactions between sampling elements (i.e., pixels) in the digital detectors Gatan uses. If you have this function, check it out in the EL/P software manual or talk to Gatan.
Could you explain briefly how I could optimize EDS acquisition? What are the things to look for? I think I tried to have the shortest data acquisition time and high beam current already. Covered above. A subtle point, perhaps, but the important parameter is probe current rather than total beam current in the microscope. Do you think 2.5 sec meter plate reading is still low? I'm really not too familiar with this.
_____________
Mo peak ... to Dr. Garber and Dr. Malcolm
I don't know about Mo aperture, but I'll check ... but I think the microscope I'm using has Be-window for EDS, is that the movable Mo-aperture you are talking about. Or is it the aperture of the microscope? (Mo) x-rays from the condenser apertures in the microscope can produce spurious signals in the EDS detector. These have no connection with EELS.
One way around the problem of molybdenum when trying to detect sulphur is to use the L2,3 edge of sulphur.
this edge is at 165 eV which is far enough away from the plasmon, so no problems there. other major in that region are
Ho N 45 161 eV Y M 45 157 Dy N 45 154 Tb N 45 147 Gd N 45 140
I hope that these are not in your sample but if they are not strong edges. To get a good signal in this energy range I would have convergence angle of around 11 mrads a COLLECTOR angle of around 3.4 mrads this is the important one
Acquisition set at 0.3 eV per channel 10s integrate time 3 times per acquire attenuator off
The probe should be 1 nm in STEM mode your VSM should have an offset of 100 eV
Sorry, but that Electrohome monitor is not a standard RGB monitor. Ours failed and I remember Kevex telling me that the scan rates were pushed a little harder to get the resolution they used on the Deltas. I tried several RGB monitors including an old Sony TV monitor, a DEC computer monitor, and new Hitachi and NEC computer monitors with RGB inputs. None of them could quite sync up.
We replaced ours with one from another Delta system being retired on campus. I had also asked the list and gotten responses from several others willing to help me out. But now, our Delta is sidelined and I might be persuaded to finally part with the whole thing. Let me know how the search goes.
Waren
At 08:45 AM 11/24/98 -0500, Kalman wrote: } } On Mon, 23 Nov 1998, Craig Franklin wrote: } } I am looking for a used color monitor for a Kevex Delta 1 EDX System. It is } } a model 38-DO5IMA-UU free standing monitor with 5 RCA plugs exiting the } } rear. } } If those plugs are labelled R,G,B,H,V, it is probably a } standard RGB monitor and there are many replacements from } many manufacturers. If it is an older machine, the } bandwidth is probably not great making replacement } relatively easy. Surely, someone in the lab/department has } a computer monitor with RGB inputs you could try out. } } Kal }
---------------------------------------------------- Warren E. Straszheim 23 Town Engineering Iowa State University Ames IA, 50011-3232
For what you are going to pay for a darkroom setup (enlarger, supplies, chemicals, paper, etc.) you could afford to go with a digital system. I bought a Polaroid SprintScan 45 for around $8100 (or much less than a good enlarger) This will do full TEM negs at 2000 dpi and 35mm at 4000 dpi. For the full TEM negative, this gives an enlargement factor of about 6.7X if you print to a 300 dpi printer. Although you should really consider getting a sublimation dye printer, you can get marvelous results from $300-$400 inkjet printer. I use an HP 892C at work and a HP 722c at home. Both of these do fantastic photo quality output when used with the Photo Deluxe Paper. -Scott
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: "740206-at-ucl.itri.org.tw"-at-Sparc5.Microscopy.Com To: Microscopy-at-Sparc5.Microscopy.Com -----------------------------------------------------------------------.
Fellow microscopists, I am setting up a darkroom for my electron microscope laboratory recently and looking up a suitable enlarger. I would appreciate your suggestions and comments on what brand/ specifications I need for my TEM, SEM and optical microscopes. Thanks Ren-Jye
I do not belong to this list, however it was suggested that I post my question here. Any help would be greatly appreciated. My e-mail address is jane-at-cc.usu.edu
My question pertains to transmission electron microscopy (TEM) on mammilian embryos. I have done a literature search and will be getting copies of articles soon. However, I was wondering if anyone out in Histoland has processed mammilian embryos for TEM? From the materials and methods of the articles I have seen, they are fixed in standard EM fixative, rinsed, fixed with OsO4, rinsed, dehydrated, and embedded in Epon - all similiar to what I am used to.
Questions: Do you use standard BEEM capsules?
I'll have a dissecting microscope to use when I am processing the samples, however, how do you see them to put them in the block?
Some of the articles I have read suggest embedding the embryos is agar during processing - has anyone done this?
Thank you in advance,
Jane
M. Jane Chambers, MS, LAT Research Technician USDA/ARS Poisonous Plant Research Lab 1150 East 1400 North Logan, UT 84341 Phone: (435) 752-2941 FAX: (435) 753-5681 jane-at-cc.usu.edu
----snip---------- } I wouldn't count on them for long term storage of critical information. we } went to a relatively cheap digital image capture system (Snappy Image } capture card on the NTSC outlet of our AMRAY 1830) and are well pleased with } the results. We typically take both thermal prints and a digital image on } all images. } } John Giles } Principal Materials Engineer } Honeywell Space Systems
"relatively cheap"???? How about DIRT CHEAP!
Seriously though, it brings up a question which I don't think I've heard addressed here even though image capture comes up frequently. I am currently using a Snappy for capturing (on a dinosaur of a computer, I might add!) from an optical scope and am planning on upgrading to a real capture card.
Has anybody compared the consumer capture cards (e.g. All-In-Wonder, for a couple hundred dollars) with the higher cost cards (e.g. Flashpoint, costing a thousand or more)?
I'd be interested in hearing thoughts from the list.
Jim Passmore Analytical Chemist Cryovac Divsion Sealed Air Corporation james.passmor-at-sealedair.com
Rather than purchase a monitor to do this, have you considered using a frame grabber to capture and display the images? You could use a standard PC monitor for viewing the images. This way, you would not only be able to visualize your images, but you'd have the added capability of capturing and storing the images. We have a couple of boards that might do the job, cost is between $1300 - $2500 for non-standard signals. If you can send me specifications for the video signal, I can tell you what you'd need. The boards are auto synching, so all you'd have to do is attach your video cables, and tell the board to determine the signal. This is a very simple procedure which does not require you to program in a bunch of timing values. Feel free to email me if you are interested.
Disclaimer: Obviously, I have a vested interest here. I can solve your problem, but in the process I am looking to sell you this equipment.
Thanks, Jim Haley
****************************** 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 ******************************
Craig Franklin 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 looking for a used color monitor for a Kevex Delta 1 EDX System. It is } a model 38-DO5IMA-UU free standing monitor with 5 RCA plugs exiting the } rear. } } If anyone can help, please contact me at 303-689-2224 or e-mail } franklin-at-idcomm.com... } } Thank you, } Craig Franklin } franklin-at-idcomm.com
We have had good luck with our Snappy card. Ours is relatively new (I think the latest version). This is the third or fourth capture board we have had in our system over the last 7-8 years.
Our initial board was a US Video VGA capture board. It was ~$700 and seemed to be oriented towards the TV industry as it had stuff for blue screens and alphanumerics. Quality was okay, not great, but it seemed to be somewhat outdated when we bought it. Chalk that one up to inexperience.
We decided the next board would be more suited towards our technical application. Our budget for this was $1-2K. We chose the Targa 64 card. The resolution was much better than the US Video, but it wasn't the most user friendly software and we had all grades of problems when we went to a Windows operating system (from DOS).
The next card was a Snappy. the resolution was as good as the Targa and the software was much more user friendly. We had this one for a while and then purchased a later version since it was so cheap (~$100).
In our application, the high end boards don't do a lot for us, since we are getting an NTSC video signal out of our AMRAY 1830. Our other option would be a retrofit with a system to capture the digital signal, but we haven't reached that step yet in either need or budget.
John Giles Principal Engineer Honeywell Space Systems
} "relatively cheap"???? How about DIRT CHEAP!
} Seriously though, it brings up a question which I don't } think I've heard addressed here even though image } capture comes up frequently. I am currently using a } Snappy for capturing (on a dinosaur of a computer, I } might add!) from an optical scope and am planning on } upgrading to a real capture card.
} Has anybody compared the consumer capture cards (e.g. } All-In-Wonder, for a couple hundred dollars) with the } higher cost cards (e.g. Flashpoint,costing a thousand or } more)?
} I'd be interested in hearing thoughts from the list.
} Jim Passmore } Analytical Chemist } Cryovac Divsion } Sealed Air Corporation } james.passmor-at-sealedair.com
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I am looking for a used color monitor for a Kevex Delta 1 EDX System. It is a model 38-DO5IMA-UU free standing monitor with 5 RCA plugs exiting the rear.
If anyone can help, please contact me at 303-689-2224 or e-mail franklin-at-idcomm.com...
by rwja.UMDNJ.EDU (8.8.6 (PHNE_14041)/8.8.6) with ESMTP id SAA16673; Tue, 24 Nov 1998 18:04:52 -0500 (EST) Message-ID: {365B64B5.80DF1B0C-at-umdnj.edu}
740206-at-ucl.itri.org.tw-at-sparc5.microscopy.com wrote: } } Fellow microscopists, } I am setting up a darkroom for my electron microscope laboratory recently and looking up a suitable enlarger. I would appreciate your suggestions and comments on what brand/ specifications I need for my TEM, SEM and optical microscopes. } Thanks } Ren-Jye
My choice would be a Beselar 45MXT enlarger, about $1000. Will do everything from 35 mm to 4 by 5 film. Be sure to get a high quality enlarging lens, such as an El Nikor or Schneider which will cost about $250-$300. A 135 mm lens is the correct focal length for 3.25 by 4 EM film. You can add other lenses to the system as you need them. With reasonable care such equipment will last forever.
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 ***************************************************************
Evex Service, a division of Evex Analytical, offers repair exchange = parts and detector upgrades for many older and some newer style x-ray = analyzers. Evex Service, maintains a full line of Kevex, Tracor, Noran, = and PGT Analyzers for its Service Contract Customers. If you need parts = or technical support, feel free to contact any of our experienced = service engineers, the average has 10+ years in x-ray microanalysis.
Peter Tarquinio Evex Analytical 857 State Road Princeton, NJ 08540
-----Original Message----- } From: Gary M. Easton {gary.easton-at-scannerscorp.com} To: MSA Listserver {microscopy-at-sparc5.microscopy.com}
Dear Colleagues:
Once again I have found myself with a few extra hotel rooms for a conference. I have 2 rooms available for the Marriott in Boston for the Materials Research Society Meeting. The rooms are available for arrival Saturday November 28 for 6 nights at the conference rate which I believe = is $128.
I made this same offer for the MSA meeting and realized that many people for one reason or another need last minute hotel rooms. If that is the case for MRS meeting, please let me know immediately. If I don't have an= y takers, I'll be canceling the rooms tomorrow night.
I am making some 200 mesh coated nickel grids for some immuno work. Normally we use Pioloform from Agar Scientific to make our support films, but I have never used coated grids for immuno work before. Does anyone have experience using Pioloform-coated grids for immuno staining?
If so will they work OK or will I have to resort to Formvar ?
{html} {!DOCTYPE HTML PUBLIC "-//W3C//DTD W3 HTML//EN"} {HTML} {HEAD}
{META content=text/html;charset=iso-8859-1 http-equiv=Content-Type} {META content='"MSHTML 4.72.3110.7"' name=GENERATOR} {/HEAD} {BODY bgColor=#ffffff} {DIV} {FONT color=#000000 size=2} Hi, {/FONT} {/DIV} {DIV} {FONT color=#000000 size=2} I'm in need of a EDS HV bias module, NIM style, with an adjustable output. I can't use the one made by Tracor as it has to be paired with their pulse processor. Maybe one out of an old KEVEX 5100 system or similar. Thanks {/FONT} {/DIV} {DIV} {FONT color=#000000 size=2} {/FONT} {/DIV} {DIV} {FONT color=#000000 size=2} {/FONT} {/DIV} {DIV} {FONT color=#000000 size=2} Gary M. Easton, Pres. {/FONT} {/DIV} {DIV} {FONT color=#000000 size=2} SCANNERS CORPORATION {/FONT} {/DIV} {DIV} {FONT color=#000000 size=2} Third Party SEM Service/Maintenance {/FONT} {/DIV} {/BODY} {/HTML}
I would like to thank all those that responded to my question. I now feel that I know more than I did before I asked (heh, imagine that...). As time permits, I may correspond with those offering their expertise or services. To the rest, thank you very much. Have a wonderful Thanksgiving Holiday!!
Best Regards,
Tim tim_wallace-at-doh.state.fl.us
Go FSU Seminoles! (oops, sorry about that.) ________________________________________________ Tim Wallace, Environmental Specialist II Florida Department of Health Volusia County Health Department, Environmental Health Division Indoor Air Assistance / Lead Monitoring Programs 501 S. Clyde Morris Blvd., Daytona Beach, Florida 32114 USA phone: 904.947.3484 / fax: 904.947.3485 http://www.state.fl.us/cf_web/D12/cphu/ehprgms/iaq.html +++++++++++++++++++++++++++++++++++++
P.S. Just in case that anyone is interested, there is a listserve on the topic of Indoor Air Quality called iaq-at-onelist.com. A subscription can be had at {http://www.onelist.com}, go to "find a list", go to "environmentalism" and dbl. click, scroll down to the IAQ list description. There you can subscribe if interested. I have no financial interest in the IAQ listserve.
Hi all Re the thread on HEPES as buffer vehicle for glut fixation:
A long time ago we did some work on the buffering capacity of some common (and a few not-so-common) buffers. HEPES was one of these. This buffer has a Pka right in the desired area (approx 7.5) and has reasonable buffering capacity. Cost is the primary disadvantage.
The reference is: Coetzee & van der Merwe (1987): Some characteristics of the buffer vehicle in glutaraldehyde-based fixatives. Journal of Microscopy, 146, 143-155.
Jan Coetzee
} I've heard that this buffer does not hold it's buffering capacity very } well. though I've no personal experience with it. } } Perhaps someone out there has worked with it.
Prof Jan Coetzee Head: Lab for Microscopy and Microanalysis Tel:+27-12-420-2075 University of Pretoria Fax:+27-12-362-5150 Pretoria 0002, South Africa http://www.up.ac.za/science/electron/emunit1.htm
I could not access email for a while due to network problems in my institute. For the question I asked about carbon rod sharpener, many people sent me good pieces of information, suggestion, or even a free rod sharpener. They were all very helpful to me. Thank you all who helped me and thank ListServer.
Especially many thanks to: Dr Laurence Tetley at University of Glasgow Olli Taikina-aho at University of Oulu, FINLAND Malcolm Haswell at University of Sunderland Dr. Ming H. Chen at University Of Alberta, Canada John Heckman, MSU Center for Electron Optics James Young David Henriks, South Bay Technology, USA Bill Tivol Markus F. Meyenhofer, Microscopy Labs, USA Paul Nolan
Jondo Yun Department of Inorganic Materials Engineering Kyungnam University 449 Weolyeong-dong Masan, 631-701 Korea 82-551-249-2697 (office) 82-551-248-5033 (fax) 82-551-249-2692 (department office) 82-551-249-2719 (laboratory) 82-551-249-2564 (EM lab) email: jdyun-at-hanma.kyungnam.ac.kr
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In response to Ren-Jye who had questions about conventional darkrooms, I have a suggestion that might make your work easier. Micrographs of metals taken on a TEM tend to have high contrast when developed in D-19, making printing difficult. I switched to Acufine Co.'s Diafine developer. It requires soaking the film 3 min. in each of two solutions, then fixing and rinsing normally. It provides CHEMICAL DODGING in effect, reducing contrast to a very printable level. It has good shelf life as mixed and is more expensive-but the time and print paper saved make up for that. I normally can print negs. on No. 2 contrast paper. Bernie Kestel {Kestel-at-anl.gov}
(1.37.109.15/16.2) id AA140123896; Wed, 25 Nov 1998 11:11:36 -0600 Posted-Date: Wed, 25 Nov 1998 11:11:36 -0600 Received-Date: Wed, 25 Nov 1998 11:11:36 -0600
A few weeks ago I posted a question about etching glycol methacrylate polymer (specifically JB4, Polysciences) for immunohistochemistry in small tissue aggregates. I'd like to thank Hank Adams for suggesting an etching technique that worked. I was able to see specific staining in my samples using his suggestion.
} Try etching for 3-5 minutes in sodium ethoxide soln 1:1 with toluene, } then go to 100% EtOH down to water, buffer, etc. } } Ethoxide Solution: } 75gm NaOH in 946 ml absolute ethanol; let stand 10-14 days or until } pellets dissolve with occasional stirring. Solution should turn brown } when ready.
} Hank Adams } Cell Biology } Integrated Microscopy Core } Baylor College of Medicine } One Baylor Plaza } Houston, Tx 77030
Susan A. Fugett
Department of Chemical Engineering and Materials Science Phone: 612-625-8803 University of Minnesota 612-625-0808 421 Washington Ave SE Fax: 612-626-7246 Minneapolis, MN 55455 Email: fugett-at-cems.umn.edu
Robert Santoianni {Robert_Santoianni-at-emory.org} wrote:
} To Harris Reavin, } The evolution of clearing solvents is interesting. Toluene (actually } chloroform) works best, but is a no-no because of carcinogenicity and } environmental (disposal) problems. Xylene has been the solvent of } choice for both clearing and dewaxing, but again, health effects and } disposal are major concerns. Limonene products came on the scene as } the "magic" xylene substitute, boasting the fact(?) that they could be } dumped down the drain. Well, in my experience they don't perform well, } our local sewer authorities don't want them in the system and you can } smell them out in the parking lot! Aliphatic hydrocarbons, such as certain } products from Richard-Allen Scientific 800-522-7270 and ANATECH } 800-ANATECH have a proven track record as xylene substitutes. Check } with these companies, you might persuade them to send you some } samples...and they both are very knowledgeable and helpful and offer all } kinds of expert advice. In my opinion, you'll get good clearing with } aliphatic hydrocarbons, but you will have to do your whole processing } procedure under a fume hood. Remember gloves, goggles and } apron...be safe! } Good luck, } Bob Santoianni } Emory University Hospital } Atlanta, GA } robert_santoianni-at-emory.org
I have recently begun using limonene as a clearing agent, and have received decent results. I am looking at cadaveric tissue that has been in phenol for a while (years), but I've been able to get decent preservation using limonene as a clearing agent. I'm also working around students who are not professionals used to working with more dangerous sovents, so I feel a little safer giving them limonene. As for odor, like many people with a biochem background, I have a weak sense of smell, so I don't really notice it.
I am aware my needs are very different than a path lab making surgical decisions based on their results, and I'm also not doing this work full time. If I was doing it day in and day out, poorer preservation and overwhelming odor might be more of an issue.
Also, I'm interested in the body of research suggesting limonene has antitumour effects. While I do not take a nip out of the bottle or intentionally inhale fumes, I sometimes wonder if it might help neutralize the "sins of my youth"(exposure to aldehydes, Osmium and what not). What are others experiences with the magic orange juice? Given it's low flash point, is there a safe way to dispose of it by burning it? I'm not suggesting this, just curious if it's been tried or thought out.
Also, does anyone know if its clearing properties deteriorates if BHA or BHT isn't added? Or what the optimal concentration of these additives is? I'm also looking into cheaper sources. I can get 55 gallons of a higher grade of limonene from a chemical company for less than half of what one of the histology companies is charging for five gallons, but I don't really feel like storing 55 gallons of a flammable in a small lab.
Charlie Ginsburg NCC Research Dept. Lombard IL
_________________________________________________________ DO YOU YAHOO!? Get your free -at-yahoo.com address at http://mail.yahoo.com
Dear List I have the opportunity to purchase an inverted fluorescent microscope,to use for viewing cells infected with a virus expressing green fluorescent protein, and to take photographs or capture an image that will let me count plaques, measure intensity of the signal etc. As this has to live in the virus room, it may not be practical to have a multipurpose computer set up alongside, so I was thinking about a digital camera with discs so users could take information away to analyse elsewhere. I am inundated with options but I wonder if anyone has such a setup that they are happy with and could recommend. My budget is =A325K absolutely max. Please respond off the list and I will summarise at a later date! Many thanks Vivien Mautner
Vivien Mautner CRC Institute for Cancer Studies The University of Birmingham Edgbaston BIRMINGHAM B15 2TA
while we can't help you with the diffractometer and it's components, we can probably help you with your CCD camera, image acquisition and processing needs. We have a camera that allows to acquire 10-bit images with extremely short or very long exposure times to cover the wide dynamic range of diffraction patterns.
If you are interested in getting more info, please contact me directly.
Michael Bode
Michael Bode, Ph.D. Soft Imaging System Corp. 1675 Carr St., #105N Lakewood, CO 80215 phone: (888) FIND SIS fax: (303) 234-9271 email: info-at-soft-imaging.com
================== Disclaimer: Soft imaging System Corp produces and sells image acquisition and processing systems. We therefore have a vested interest in some of the items mentioned above. ==================
--------- Forwarded Message ---------
DATE: Mon, 23 Nov 1998 08:02:32 } From: "Dennis C. Winkler" {Dennis_Winkler-at-nih.gov} To: Microscopy-at-Sparc5.Microscopy.Com
------------------------------------------------------------------------ The Microscopy ListServer -- Sponsor: The Microscopy Society of America
Hello all,
We are planning to rebuild the optical diffractometer we use to assess the quality of our TEM micrographs. Anyone have a favorite design or modification for an optical diffractometer (and perhaps an associated reference)? We are looking to build a vertical set up similar to the one in:
We'd also like to add a CCD camera with a high quality thermal printer.
Any recommendations?
Thanks,
- Dennis
------------------------------------------------------------------------ ------ Dennis C. Winkler, PhD. Phone: (301) 496-0131 Laboratory of Structural Biology Research Fax: (301) 480-7629 NIAMS, National Institutes of Health Email: Dennis_Winkler-at-nih.gov Bldg. 6, Room B2-26, MSC-2717 Bethesda, MD 20892-2717, U.S.A.
--------- End Forwarded Message ---------
-----== Sent via Deja News, The Discussion Network ==----- http://www.dejanews.com/ Easy access to 50,000+ discussion forums
I've replaced our Targa framegrabber with the Snappy ($3000 vs. $100) and am much happier with the Snappy. Simple to setup and simple to use. Trying to figure out the Targa manual set (yes, set!) is no picnic. The Snappy manual alone is worth the price, simply for entertainment value. Even their licensing agreement is hilarious. For standard NTSC resolution cameras, I wouldn't bother with anything else.
} ------------------------------------------------------------------------ } 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 } -----------------------------------------------------------------------. } } ----snip---------- } } I wouldn't count on them for long term storage of critical information. we } } went to a relatively cheap digital image capture system (Snappy Image } } capture card on the NTSC outlet of our AMRAY 1830) and are well pleased with } } the results. We typically take both thermal prints and a digital image on } } all images. } } } } John Giles } } Principal Materials Engineer } } Honeywell Space Systems } } "relatively cheap"???? How about DIRT CHEAP! } } Seriously though, it brings up a question which I don't think I've heard } addressed here even though image capture comes up frequently. I am } currently using a Snappy for capturing (on a dinosaur of a computer, I } might add!) from an optical scope and am planning on upgrading to } a real capture card. } } Has anybody compared the consumer capture cards (e.g. All-In-Wonder, } for a couple hundred dollars) with the higher cost cards (e.g. Flashpoint, } costing a thousand or more)? } } I'd be interested in hearing thoughts from the list. } } Jim Passmore } Analytical Chemist } Cryovac Divsion } Sealed Air Corporation } james.passmor-at-sealedair.com
--
James M. Ehrman Digital Microscopy Facility Mount Allison University Sackville, NB E4L 1G7 CANADA
A year or so ago the address for Taab Labs Equip. Ltd. was 3 Minerva House, Calleva Industrial Park, Aldermaston, Berkshire RG7 4QW, UK, Tel: 0734-817775, Fax: 0734-817881
Wilbur C. Bigelow, Prof. Emeritus Materials Sci. & Engr., University of Michigan Ann Arbor, MI 48109-2136 e-mail: bigelow-at-umich.edu; Fx:734-763-4788; Ph:734-764-3321
I could not access email for a while due to network problems in my institute. For the question I asked about carbon rod sharpener, many people sent me good pieces of information, suggestion, or even a free rod sharpener. They were all very helpful to me. Thank you all who helped me and thank ListServer.
Especially many thanks to: Dr Laurence Tetley at University of Glasgow Olli Taikina-aho at University of Oulu, FINLAND Malcolm Haswell at University of Sunderland Dr. Ming H. Chen at University Of Alberta, Canada John Heckman, MSU Center for Electron Optics James Young David Henriks, South Bay Technology, USA Bill Tivol Markus F. Meyenhofer, Microscopy Labs, USA Paul Nolan
Jondo Yun Department of Inorganic Materials Engineering Kyungnam University 449 Weolyeong-dong Masan, 631-701 Korea 82-551-249-2697 (office) 82-551-248-5033 (fax) 82-551-249-2692 (department office) 82-551-249-2719 (laboratory) 82-551-249-2564 (EM lab) email: jdyun-at-hanma.kyungnam.ac.kr
Our group currently has a JEOL FX 2000 TEM available for sale to anyone interested. The system includes a cold and hot stage, PC driven X-ray system,and STEM attachment. Please respond if interested.
M.W. Rigler, Ph.D. MAS, Inc. Suwanee GA 770-866-3218
I am currently in the process of making cross-section samples of a machined Ni-Cr alloy. The samples are first cleaned in NaOH (& Al. clip) and rinsed in warmed distilled H2O. They are then ultrasonically cleaned in distilled acetone, and also suspended in an acetone reflux vapor degreasing for 20 minutes.
I immerse the samples in the Nickel "Electroless" plating solution for ~20 minutes at 90-100 C. Fine bubbles form indicating that plating is proceeding but when I remove the sample, a discolouration appears on the surface and the film doesn't appear to adhere very well.
Any suggestions?
I eventually want to Nickel electroplate to about 2-3 mm. then prepare the cross-sections.
thanks in advance
Fred
******************************************************** Fred Pearson Brockhouse Institute for Materials Research McMaster University 1280 Main St. West Hamilton, Ontario Canada L8S 4M1
I don't know if there is anything commercially available for this for light microscopes, but you might want to check with Vince Carlino at VCR, Inc. (VCRvince-at-aol.com) Their D500i dimpler has a non-contacting sensor that has 1 um resolution. This system does have a feedback loop. He might be able to help you or direct you to someone that can. -Scott Walck
---------- } From: Bruce Brinson To: MSA Listserver -----------------------------------------------------------------------.
Hello photon fans, I need an close loop, contactless system that can check & correct the working distance between objective & glass plate to within 1u. I would appreciate any feed back this group can provide. Please contact me directly if your company has markets this type of product.
Dear sir, It will be grateful if you let me know if you have an available second hand TEM. My Fax is 009821 8895980 or 009821 2548220 Tank you very much. Hmid. habibi hrhb4-at-nrcgeb.ac.ir
The Materials Research Science and Engineering Center at the University of Houston has an opening for a research faculty member or postdoctoral fellow to manage and conduct research in their newly completed JEOL 2010F FEG-TEM facility. The facility became operational in July 1998 and consists of a JEOL 2010F FEG-TEM (high-resolution pole piece, 1.9=C5 point-to-point resolution verified), with Gatan 794 Multi-scan CCD Camera, Gatan 666 PEELS, and X-ray EDS system to be purchased in early '99. An additional JEOL 2000FX analytical TEM is available as part of shared facilities with the Texas Center for Superconductivity, together with complete sample preparation facilities.
MRSEC-UH is an NSF-funded center dedicated to research on advanced oxides for solid-state ionics applications and a background in TEM-based oxide structural chracterization is desirable but not essential. The successful candidate will conduct cooperative research related to the various active projects in the Center, and will manage the day-to-day use of the facility. Development of the facility's full potential in the area of HRTEM imaging and nano-probe microanalysis will be expected.
Interested individuals should respond (off-line please!) to:
Professor Allan J. Jacobson, Director MRSEC-UH Department of Chemistry University of Houston, Houston, TX 77204 ajjacob-at-uh.edu
Additional information regarding MRSEC-UH can be found at: www.uh.edu/mrsec. The University of Houston is an Equal Opportunity/Affirmative Action Employer. Minorities, women, veterans and persons with disabilities are encouraged to apply.
We are faced with replacing an ancient Haskris on a diff-pumped Balzer 301. Wondering whether Haskris is still the way to go, and whether there are reliable alternatives. We are working with a water-cooled compressor...Thanks...Tom Reese
Dear driver experts, Our laboratory have a old monitor Sony, model GDM 1952 Trinitron. What kind of graphic cart and driver we can use for our windows applications Many thanks OPEA 114, rue de la Jarry 94300-VINCENNES (FRANCE) e-mail: opea.hoan-at-wanadoo.fr
I'm studying arthropod (mostly small crustacean) appendages. Some of the mounts are relatively thick, with fine structures well below the coverslip. I've read Barron's fine book and noted his comments on thick mounts causing spherical aberration. I have considered modifying my scope to have adjustable tubelength. But it occurred to me that some correction could be obtained by using a high index mounting medium, i.e., higher than immersion oil / coverslip... and higher than chitin to get contrast. Cargille's "Meltmount" sounds good but I prefer a water soluble medium. Questions follow. --- Am I correct in my belief that a high index medium will nullify some spherical aberration in these thick mounts? --- One type of "high index" medium uses Chloral Hydrate, which is hard to obtain and I doubt the index is very high. I've heard recommendations of heavy halogen salts... any recipes will be appreciated. --- I've been using mostly gum - glycerin - water. Another problem there is shrinkage with distortion of the coverslip, which can't be good for resolution. Any hints to minimize that will be appreciated. --- The above medium is a colloid(?); am I losing contrast due to light scatter? Would a true solution be *significantly* better with brightfield? Darkfield? --- Does anyone know the refractive index of chitin? (Looks barely above imm. oil). How much higher should the medium index be to get good contrast AND resolution... without too much Becke line / white line?
I hope no one minds these low - tech questions on a high - tech newsgroup. I've done a fair amount of web-search without finding the information I'm looking for.
I shall be on leave during 3-20 Dec. I will switch on the auto-answer function (every coming e-mail will get auto-answer from my computer) and I do not want this auto-answer affect every member of this mail list. So I want to unsubscribe this mail list. Please take my name from the mail list. Thank you very much.
-----Original Message----- From: malcolm.haswell-at-sunderland.ac.uk [SMTP:malcolm.haswell-at-sunderland.ac.uk] Sent: Friday, November 13, 1998 1:21 AM To: Subject: Re: To CPD or to HMDS, that is the ques
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Could something be sent to this list too please - it would certainly
interest me.
thanks
Malcolm Haswell Electron Microscopy School of Health Sciences Fleming Building University of Sunderland SUNDERLAND SR1 3SD Tyne and Wear UK
Tel (0191) 515 2872 e-mail: malcolm.haswell-at-sunderland.ac.uk ---------- } From: ROBIN CROSS To: Paula Sicurello Cc: microscopy Subject: Re: To CPD or to HMDS, that is the ques Date: 12 November 1998 15:51
Hello Paula
} Or I was considering HMDS, has anybody out there used HMDS for } things as delicate as cells?
Shirley Pinchuck, in this lab, has done a fair amount of work using HMDS, including comparisons of HMDS with other methods such as CPD, cryo-SEM, etc. I will ask her to fax you her protocols as well as an abstract of a conference presentation on some of the comparative work.
I hope this helps.
Regards
Robin
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
} -----Original Message----- } From: ZHANG Tiejun } Sent: Monday, November 30, 1998 6:11 PM } To: DAI Jiyan; DAI Jiyan } Subject: EM survey } } Dear all: } I post this message for a TEM service engineering. } } Please kindly let me know which EM Lab has 3 sets of TEM made by JEOL. } Thank you in advance!
Dear driver experts, Our laboratory have a old monitor Sony, model GDM-1952 Trinitron with input RGB, H.synchro. and V.synchro. What kind of graphic card and driver we can use for our windows applications Many thanks Hoan OPEA 114, rue de la Jarry 94300-VINCENNES (FRANCE) e-mail: opea.hoan-at-wanadoo.fr
I suppose that you could use the monitor on most any card with an RGB output. However, I don't know how many windows applications there would be that would make use of it. It might be that frame grabber/display cards could make use of it in connection with image analysis programs. Matrox and other companies could probably answer those questions for you and let you know what products they have that would drive that monitor. Their cards would likely come with the necessary drivers.
I am not familiar with the size of that monitor or its resolution specifications. If you are thinking of using it as your main monitor, you may well be able to find or make an adapter cable to go from a standard VGA connector to the BNC connectors you probably need for RGB. However, there would likely be issues of the monitor being able to run at the scan frequencies that VGA cards support. The resolution may also be unsatisfactory. Very old VGA monitors specified a dot pitch of 0.39 mm. Then came monitors with 0.28 mm pitch and now 0.22 mm. If your Sony does not have such a fine pitch, you would probably find the graphics blurred.
That's about as much as I can tell you.
At 01:34 PM 11/30/98 +0100, you wrote: } } Dear driver experts, } Our laboratory have a old monitor Sony, model GDM-1952 Trinitron with } input RGB, H.synchro. and V.synchro. What kind of graphic card and } driver we can use for our windows applications } Many thanks } Hoan } OPEA } 114, rue de la Jarry } 94300-VINCENNES (FRANCE) } e-mail: opea.hoan-at-wanadoo.fr
It is true that, for large angles of incidence, the spherical aberration of a plate is proportional to t/N (thickness over refractive index). So higher index might just give you a better image, if the other aberrations don't creep up on you, particularly longitudinal chromatic. But wouldn't you want a higher index cover slip as well?
best regards mark
Mark W. Lund, PhD VP Engineering } } 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
"This is a YOUNG business...How can I tell you what YOUR job is when I don't know what MINE is?" --Pogo
Hello all;
I'm studying arthropod (mostly small crustacean) appendages. Some of the mounts are relatively thick, with fine structures well below the coverslip. I've read Barron's fine book and noted his comments on thick mounts causing spherical aberration. I have considered modifying my scope to have adjustable tubelength. But it occurred to me that some correction could be obtained by using a high index mounting medium, i.e., higher than immersion oil / coverslip... and higher than chitin to get contrast. Cargille's "Meltmount" sounds good but I prefer a water soluble medium. Questions follow. --- Am I correct in my belief that a high index medium will nullify some spherical aberration in these thick mounts? --- One type of "high index" medium uses Chloral Hydrate, which is hard to obtain and I doubt the index is very high. I've heard recommendations of heavy halogen salts... any recipes will be appreciated. --- I've been using mostly gum - glycerin - water. Another problem there is shrinkage with distortion of the coverslip, which can't be good for resolution. Any hints to minimize that will be appreciated. --- The above medium is a colloid(?); am I losing contrast due to light scatter? Would a true solution be *significantly* better with brightfield? Darkfield? --- Does anyone know the refractive index of chitin? (Looks barely above imm. oil). How much higher should the medium index be to get good contrast AND resolution... without too much Becke line / white line?
I hope no one minds these low - tech questions on a high - tech newsgroup. I've done a fair amount of web-search without finding the information I'm looking for.
I was wondering if it would be easier to find an objective with a correction collar on it? We have a 40x with a collar that we use for all our samples that are outside the standard thickness. It helps a lot. But I don't know if it would give enough resolution for your situation.
Bob Derm Imaging Center U of W
On Sun, 29 Nov 1998 Sjoplinh-at-aol.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 } -----------------------------------------------------------------------. } } } Hello all; } } I'm studying arthropod (mostly small crustacean) appendages. Some of the } mounts are relatively thick, with fine structures well below the coverslip. } I've read Barron's fine book and noted his comments on thick mounts causing } spherical aberration. } I have considered modifying my scope to have adjustable tubelength. But } it occurred to me that some correction could be obtained by using a high index } mounting medium, i.e., higher than immersion oil / coverslip... and higher } than chitin to get contrast. Cargille's "Meltmount" sounds good but I prefer } a water soluble medium. Questions follow. } --- Am I correct in my belief that a high index medium will nullify some } spherical aberration in these thick mounts? } --- One type of "high index" medium uses Chloral Hydrate, which is hard to } obtain and I doubt the index is very high. I've heard recommendations of } heavy halogen salts... any recipes will be appreciated. } --- I've been using mostly gum - glycerin - water. Another problem there is } shrinkage with distortion of the coverslip, which can't be good for } resolution. Any hints to minimize that will be appreciated. } --- The above medium is a colloid(?); am I losing contrast due to light } scatter? Would a true solution be *significantly* better with brightfield? } Darkfield? } --- Does anyone know the refractive index of chitin? (Looks barely above } imm. oil). How much higher should the medium index be to get good contrast AND } resolution... without too much Becke line / white line? } } I hope no one minds these low - tech questions on a high - tech newsgroup. } I've done a fair amount of web-search without finding the information I'm } looking for. } } Regards, } } Scott Harden sjoplinh-at-aol.com } } }
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11/30/98 9:00 AM
A friend of mine working on a similar nickel plating problem years ago did not have good luck with electroless nickel plating-it did not adhere well. He made test runs with electroplating only on scrap material first. The general procedure was to have the specimen immersed in the plating bath, start with a low current and voltage in REVERSED polarity mode first for a few seconds, then change to the plating mode polarity and plate for several hours at a slow rate for good adhesion and fine grain,(again at a low voltage/current setting). This process is called the "strike" in library books. Then a higher voltage and plating rate may be used to build up a useful nickel plating thickness. This sequence resulted in good adhesion to the specimen. The Ph of the solution had to be kept near some optimum value. A bag of nylon polishing cloth was tied around the sacrificial electrode to keep residues from moving to the specimen being plated. Reading up on the process shoul! d provide the needed details.
Bernie Kestel {kestel-at-anl.com}
--------------------------------------
Greetings:
I am currently in the process of making cross-section samples of a machined Ni-Cr alloy. The samples are first cleaned in NaOH (& Al. clip) and rinsed in warmed distilled H2O. They are then ultrasonically cleaned in distilled acetone, and also suspended in an acetone reflux vapor degreasing for 20 minutes.
I immerse the samples in the Nickel "Electroless" plating solution for ~20 minutes at 90-100 C. Fine bubbles form indicating that plating is proceeding but when I remove the sample, a discolouration appears on the surface and the film doesn't appear to adhere very well.
Any suggestions?
I eventually want to Nickel electroplate to about 2-3 mm. then prepare the cross-sections.
thanks in advance
Fred
******************************************************** Fred Pearson Brockhouse Institute for Materials Research McMaster University 1280 Main St. West Hamilton, Ontario Canada L8S 4M1
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Replying to Barry Shaw's question about using Pioloform-coated grids---Pioloform will do just fine in immuno procedures. As with Formvar, you will see some of the same problems with electrostatic binding of immunogold probes to the film, but they may be prevented by incorporating a surfactant such as Tween-20 (0.05-0.1%) to your dilution and wash buffers or by using a high salt concentration in the wash buffers after the incubation in immunogold. If you are using the Pioloform to support sections instead of cells or cell fractions, the background on the Pioloform film alone may not interfere, and as long as the sections are free of background, you may not need to use a surfactant or otherwise modify the washing procedure.
Vicky
Victoria J. Madden Microscopy Services Laboratory Dept. of Pathology and Laboratory Medicine University of North Carolina-Chapel Hill vmadden-at-med.unc.edu
(SMTPD32-4.06) id AA69A80009A; Mon, 30 Nov 1998 11:40:09 EDT Message-Id: {3.0.3.32.19981130115108.0077ae70-at-mail.map.com} X-Sender: mme-at-mail.map.com X-Mailer: QUALCOMM Windows Eudora Pro Version 3.0.3 (32)
Dear Scott,
You have asked a series of interesting questions.
First, the whole contrast problem is derived from the similarity of the refractive indices of chitin and your mounting material. I did some work on this issue many years ago and offer the following solutions:
1. First: an observation:
I agree with you that the refractive index of chitin is about the same as immersion oil (1.515). As I remember, we tried immersion oil as a mounting material and the chitin all but disappeared.
2 Re: refractive index changes
You don't have to have much of a refractive index change to increase the contrast. You can either go up or go down. Cargille has a whole slew of mounting oils from which to choose. You also mentioned that you would like something water soluble. How about just water (ri = 1.33)? You may be able to do a more permanent mount by making sure that the drop of water does not exceed the edge of the coverslip then sealing with something like nail polish.
3. Re:spherical aberration. The microscope's optics expect to see very specific components in the sample prep "sandwich". All the changes which cause spherical aberration (amoung other things) are derived from Snell's law (see any basic physics book). A higher refractive index will acutally accentuate the problem, not solve it. The first thing to do to optimize imaging is to check the barrel of the objectives you are using and make sure that they say "0.17" for coverslip thickness (to order: a number 1-1/2 is the equivalent). Anything thicker will contribute significantly to spherical aberration. Secondly, if you begin to get into the higher ri mounting materials, you may want to try an objective with a "coverslip correction collar". Adjusting the collar may reduce the spherical aberration problem.
3. Colloids are, indeed, serious scatterers. The resulting scatter will create glare and haze which obscures information.
4. I don't understand the "shrinkage with the coverslip" issue. Are you talking about the size of the image or a change in the prep over time?
For more specifics on coverslips, mounting media, and spherical aberration, may we suggest the book "Optimizing Light Microscopy for Biological and Clinical Laboratories"? Details are available at our website: { {http://www.MME-Microscopy.com/education}
If any of you can help this young lady, please respond to her directly, as she does not have access to the listserve.
Best regards, Steven Slap, Energy Beam Sciences
Dear all,
I am resending this message because of a "nondelivery" warning on the first try. Please ignore if you have already received it.
Thanks,
Barbara Foster
Dear Scott,
You have asked a series of interesting questions.
First, the whole contrast problem is derived from the similarity of the refractive indices of chitin and your mounting material. I did some work on this issue many years ago and offer the following solutions:
1. First: an observation:
I agree with you that the refractive index of chitin is about the same as immersion oil (1.515). As I remember, we tried immersion oil as a mounting material and the chitin all but disappeared.
2 Re: refractive index changes
You don't have to have much of a refractive index change to increase the contrast. You can either go up or go down. Cargille has a whole slew of mounting oils from which to choose. You also mentioned that you would like something water soluble. How about just water (ri = 1.33)? You may be able to do a more permanent mount by making sure that the drop of water does not exceed the edge of the coverslip then sealing with something like nail polish.
3. Re:spherical aberration. The microscope's optics expect to see very specific components in the sample prep "sandwich". All the changes which cause spherical aberration (amoung other things) are derived from Snell's law (see any basic physics book). A higher refractive index will acutally accentuate the problem, not solve it. The first thing to do to optimize imaging is to check the barrel of the objectives you are using and make sure that they say "0.17" for coverslip thickness (to order: a number 1-1/2 is the equivalent). Anything thicker will contribute significantly to spherical aberration. Secondly, if you begin to get into the higher ri mounting materials, you may want to try an objective with a "coverslip correction collar". Adjusting the collar may reduce the spherical aberration problem.
3. Colloids are, indeed, serious scatterers. The resulting scatter will create glare and haze which obscures information.
4. I don't understand the "shrinkage with the coverslip" issue. Are you talking about the size of the image or a change in the prep over time?
For more specifics on coverslips, mounting media, and spherical aberration, may we suggest the book "Optimizing Light Microscopy for Biological and Clinical Laboratories"? Details are available at our website: { {http://www.MME-Microscopy.com/education}
Chitin should have an index close to cellulose, which is listed as 1.55 (Merck). Chloral is a controlled substance. I have seen references to use of KI in water, water--glycerol, and a commercial resin but no recipes. 40% KI in water only has n 1.4027. HgI2 can be added if you can live with that (will check reference at home).
There must be polymers with minimum light scattering. I have used surgical jelly, but it contains some junk, which can be removed with a membrane filter, as a result of which the viscosity changes.
Leonard Corwin Research Chemist Fort Dodge Animal Health Princeton, NJ 08543-0400
In a message dated 98-11-30 12:08:07 EST, Barbara write:
{ {... 3. Re:spherical aberration. The microscope's optics expect to see very specific components in the sample prep "sandwich". All the changes which cause spherical aberration (amoung other things) are derived from Snell's law (see any basic physics book). A higher refractive index will acutally accentuate the problem, not solve it. { {
(Snell's sounds familiar... maybe like Barron's "sine law".) I got a headache trying to figure which would correct for a thick mount: high or low index. It seems simpler to imagine a perfect image at the eyepiece field stop, and that *projected* back to the object, which is too far below the coverslip. With a low or standard index the marginal rays converge above the object; a high index refracts them more (toward the "normal") so they should converge closer to, if not at, the object. But, I'm not at all certain of that!
} } Secondly, if you begin to get into the higher ri mounting materials, you may want to try an objective with a "coverslip correction collar". Adjusting the collar may reduce the spherical aberration problem. { {
I doubt I'll ever get that type objective, but I may modify my scope to have adjustable tubelength, which apparently does +/- the same thing. Easy on my scope as it's a monocular.
} } 3. Colloids are, indeed, serious scatterers. The resulting scatter will create glare and haze which obscures information. { {
I was afraid of that.
} } 4. I don't understand the "shrinkage with the coverslip" issue. Are you talking about the size of the image or a change in the prep over time? { {
As these mounts dry, the coverslip (std. thickness) is "mounded" (convex) over the object. This is easy to see, e.g., looking at the reflected image of a lamp (naked eye). This wouldn't matter with immersion and a n = 1.51 mount medium. With dry objectives it would seem a disaster --- the coverslip has become an unwelcome lens. I assume "Meltmounts" (and similar) are a "total solids" medium and will not shrink... but I've been hoping to avoid all the associated hassles of non- water miscible media.
} } For more specifics on coverslips, mounting media, and spherical aberration, may we suggest the book "Optimizing Light Microscopy for Biological and Clinical Laboratories"? Details are available at our website: { {http://www.MME-Microscopy.com/education}
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Many people expressed an interest in my recent posting regarding = immunocytochemical methods. For an intense immersion in = immunocytochemical techniques (cryosectioning, resins, LM, EM, stereology) = may I recommend a FEBS-sponsored course to be held in Oslo in June 1999. = Details can be found at: {http://www.hei.org/htm/curs.htm}
Regards,
Paul Webster, Ph.D House Ear Institute 2100 West Third Street Los Angeles, CA 90057 phone:213 273 8026 fax: 213 413 6739 e-mail: pwebster-at-hei.org http://www.hei.org/htm/apw.htm
by imo14.mx.aol.com (IMOv16.10) id 3GVKa26092; Mon, 30 Nov 1998 14:56:33 -0500 (EST) Message-ID: {3be829a0.3662f871-at-aol.com}
I wrote, in response to Barbara Foster's helpful information;
{ { It seems simpler to imagine a perfect image at the eyepiece field stop, and that *projected* back to the object, which is too far below the coverslip. With a low or standard index the marginal rays converge above the object; a high index refracts them more (toward the "normal") so they should converge closer to, if not at, the object. But, I'm not at all certain of that! } }
Well, I'm pretty certain that I was wrong and Barbara is right! Sorry about that and the waste of bandwidth. I think I see what I neglected to consider... embarrassing.
So I'll be using low index media for the thick mounts. That's good and bad news: The bad news is, I probably can't do much better than glycerin (n ~ 1.46) for resolution. The good news is, these things look pretty sharp in glycerin, both resolution and contrast - wise. And, low index water soluble media are common.
} From Barron: The "sine law", non-fulfillment of which leads to spherical aberration and/or coma: n R (sinU) = n' I (sin e)
I could figure out the average thickness of my "too thick" mounts and accordingly prepare a medium with the optimum index to minimize SA. Problem is, the first "n" in the above equation represents the index of the medium (media!) between object and objective. I'll have up to 3 different indices of varying thicknesses (medium - glass - air)... I doubt one could use a weighted average... this won't be easy.
Dear Listservers: I know this topic has been hashed and rehashed in this forum. But the scanner business seems to be changing very quickly. I an interested in the best scanner in the $2000-3000 range for digitizing TEM negatives. I am working with materials so I would like a scanner that has sensitivity over a large contrast range. Any recent experiences would be highly appreciated.
Unfortunately several scanners in that price range that were suggested on the listserver have been discontinued (including the highly recommended UMAX). Thanks in advance.
Does any one has a used air-cooled water recirculator (for JEOL 100CX TEM) that you are willing to part with? We recently got a used TEM but it does not come with a water recirculator.
Sincerely, Ching-Hwa Kiang
-- ============================================================================== Dr. Ching-Hwa Kiang Visiting Assistant Professor Phone: (310) 206-0563 (Office) Cram Teacher-Scholar (310) 794-4124 (Lab) Department of Chemistry and Biochemistry (310) 206-4038 (Fax) University of California chk-at-chem.ucla.edu Los Angeles, CA 90095-1569 http://www.chem.ucla.edu/dept/Faculty/ ==============================================================================
Hello, we are doing some research on protective shields for x-ray radiation. does anyone know what is the permissible or background radiation intensity? also on refering to an old publication by the British Journal of Medicine, i found out that the maximum permissible dose(MPD) for exposure of the whole body to radiation expressed in rontgens is 0.3 per week, has this level changed or is it still the same? to calculate the MPD, we need to know the dose rate (in r min), does anyone have some info on this?
it would be great if you could give me some info on these topics.
Hello, we are doing some research on protective shields for x-ray radiation. does anyone know what is the permissible or background radiation intensity? also on refering to an old publication by the British Journal of Medicine, i found out that the maximum permissible dose(MPD) for exposure of the whole body to radiation expressed in rontgens is 0.3 per week, has this level changed or is it still the same? to calculate the MPD, we need to know the dose rate (in r min), does anyone have some info on this?
it would be great if you could give me some info on these topics.
I would like to know if anyone has an endmember calculation program for WDS } analysis of garnet. If so, how is the ferric and ferrous iron problem } resolved. My instrument setup is a 733 JEOL Microprobe with 5600/5500 } software. The program need not be an on line calculation, as I have the } setup in place to transferr the probe data to a PC for manipulation with a } Lotus spreadsheet. Hope to hear from someone soon.
Hi There! I've got an announcement about our latest workshop, so here it comes. Beware this is a long message ;-)
The UC Berkeley EM Lab, Ted Pella, Inc., and Leica present:
The 3rd Annual UC Berkeley Microwave Processing Techniques Workshop January 6-8, 1999
Breakthroughs in Microwave Processing: Techniques for Light and Electron Microscopy
Date and Times: January 6-8, 1999 9:00 am to 4:30 pm each day
Instructors: Rick Giberson, Ted Pella, Inc. Dr. Kent McDonald, Supervisor UC Berkeley EM Lab
Techinques covered:
1) Immunolabelling of microwave processed tissue for LM and EM.
2) Microwave-assisted immunolabelling on plastic, paraffin and cryoultramicrotomy sections.
3) Formalin fixation in the microwave-method and theory
4) Microwave-assisted processing of fresh tissue into paraffin
5) NEW microwave processing techniques for EM and LM
Who Should Attend:
Anyone interested in improving productivity in the lab. The course will examine the latest techniques in tissue processing for electron microscopy, formalin fixation and paraffin embedding of tissue using microwave technology. The course is designed for those individuals who want to solve problems and are receptive to new ideas that work. The theory and technique of microwave processing will be examined in detail.
Location:
Electron Microscopy Lab, 26 Giannini Hall, University of California, Berkeley
Workshop Particulars: Class size is limited to 15 participants
Cost: $400.00, which includes lunch every day and one group dinner
Type: Hands-on processing. You get to embed samples with the microwave ovens.
Contacts: Kathy Stangenberg or Rick Giberson at Ted Pella, Inc. Phone: 800-237-3526 Fax: 530-243-3761 E-mail: tedpel-at-aol.com
Hotel Accomodations:
A block of rooms at the Hotel Durant has been set aside, under the name "Ted Pella, Inc.", for attendees of the workshop, although you are not required to stay there. The hotel is within walking distance to the university. Individuals must reserve and pay all charges for their room. The single rate is $99 and the double rate is $110 with a cutt off date of 12/10/98. Check-in: 3:00 pm, check-out: 12:00 pm
For Reservations: Hotel Durant 2600 Durant Ave., Berkeley, CA 94704 Phone: 510-845-8981 or 800-238-7268 Fax: 510-486-8336 E-mail: durant-at-sfa.com
For a registration form see the Ted Pella, Inc., website at www.tedpella.com or phone the contact people at Ted Pella, Inc.
Well, there you go kids, your opportunity to see what I look like after reading all of my e-mails.
Hope to see you in January!
Paula :-)
p.s. I apologize for any typos I may have done. Check out the Ted Pella website, I think they have all the info I just wrote but in a better format.
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
I finally got on the list: my original question was posted before I was "officially" on. I apologize for any responses I may have missed yesterday or this morning.
Mark wrote: } } It is true that, for large angles of incidence, the spherical aberration of a plate is proportional to t/N (thickness over refractive index). So higher index might just give you a better image, if the other aberrations don't creep up on you, particularly longitudinal chromatic. But wouldn't you want a higher index cover slip as well? { {
Well, it now looks to me like Barbara was correct (no surprise) and that, for thick mounts, a low index would help; a high index would make it worse.( I sent a "correction on correction" post earlier today but it's not here somehow.) I had a heck of a time "ray - tracing" the situation, but I think that must be correct. Interesting idea about the higher index coverslip...
Bob wrote: } } I was wondering if it would be easier to find an objective with a correction collar on it? We have a 40x with a collar that we use for all our samples that are outside the standard thickness. It helps a lot. But I don't know if it would give enough resolution for your situation. { {
A correction collared objective would be nice. Maybe someday. But if adjustable tubelength does the same thing, then I'll have correction for any old objective. You folks with your fancy bino scopes ;-) would have a hard time doing that, although there should be other optical / mechanical methods of getting the same result.
Now that a low index medium seems to be what I need, does anyone have any info on; --- Glycerin Borate; once commercially available as "Aqua - Resin". Recommended by a well known expert. Low index by itself. I'm wondering if this was just a solution of Borax (Sodium Borate?) in glycerin or if it is a compound / molecule.
---- "Water Glass", a.k.a. Sodium Silicate. Essentially a water soluble "glass". Sounds like it might be moderately low n; maybe something could be added to further lower it if it's otherwise suitable.
best,
Scott Harden
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